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 Previous Article | Table of Contents | Next Article 
Blood, Vol. 94 No. 5 (September 1), 1999:
pp. 1673-1682
Induction of Decay-Accelerating Factor by Cytokines or the
Membrane-Attack Complex Protects Vascular Endothelial Cells Against
Complement Deposition
By
Justin C. Mason,
Helen Yarwood,
Katharine Sugars,
B. Paul Morgan,
Kevin A. Davies, and
Dorian O. Haskard
From the British Heart Foundation (BHF) Cardiovascular Medicine Unit,
National Heart and Lung Institute, and the Rheumatology Unit, Division
of Medicine, Imperial College School of Technology and Medicine,
Hammersmith Hospital, London; and the Department of Medical
Biochemistry, University of Wales College of Medicine, Cardiff, UK.
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ABSTRACT |
Vascular endothelium is continuously exposed to complement-mediated
challenge, and this is enhanced during inflammation. Although the
complement-regulatory proteins decay-accelerating factor (DAF), CD59,
and membrane cofactor protein (MCP) protect endothelial cells (ECs)
against complement-mediated injury, the control of their expression and
relative contributions to vascular protection is unclear. We explored
the hypothesis that mechanisms exist which induce upregulation of
complement-regulatory proteins on ECs to maintain vascular function in
inflammation. Tumor necrosis factor alpha (TNF ) and interferon gamma
(IFN ) each increased DAF expression but not CD59 or MCP expression,
and a combination of these cytokines was more potent than either alone.
Cytokine-induced expression depended on increased DAF mRNA and de novo
protein synthesis and was maximal by 72 hours. In addition, assembly of
the membrane-attack complex (MAC) on ECs induced a 3-fold increase in
DAF expression, and this was enhanced by cytokines. DAF upregulation
was not inhibited by protein kinase C (PKC) antagonists. The increase
in DAF was functionally relevant since it reduced complement 3 (C3)
deposition by 40%, and this was inhibited by an anti-DAF monoclonal
antibody. These observations indicate that upregulation of DAF
expression by cytokines or MAC may represent an important feedback
mechanism to maintain the integrity of the microvasculature during
subacute and chronic inflammatory processes involving complement activation.
© 1999 by The American Society of Hematology.
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INTRODUCTION |
COMPLEMENT is a powerful cytotoxic
defense system which, by the nature of its effects, represents a
potential danger to host cells.1 As a consequence of this,
a complex array of protective mechanisms have evolved, including the
cell-surface proteins decay-accelerating factor ([DAF] CD55),
protectin (CD59), and membrane cofactor protein ([MCP] CD46). The
genes encoding DAF and MCP are closely related and clustered on the
long arm of chromosome 1, band 1q32, while the gene for CD59 is located on chromosome 11.2 The mechanisms by which these factors
regulate complement are distinct. DAF, which prevents the formation and accelerates the decay of complement 3 (C3) convertases,3
and MCP, which binds C3b and C4b and facilitates their degradation by
factor I,4 operate at the proximal end of the cascade,
while CD59 serves to inhibit the membrane-attack complex ([MAC]
C5b-9).5
Vascular endothelium, by virtue of its anatomic location at the
blood/tissue interface, may be directly exposed to autologous complement activation, particularly during inflammation.1
In certain instances such as severe vasculitis and, most notably, antibody-mediated hyperacute allograft rejection, this is overwhelming and results in endothelial cell (EC) death.6 However, in a variety of chronic inflammatory diseases such as atherosclerosis and
rheumatoid arthritis and systemic inflammatory states such as systemic
lupus erythematosus, endothelium is continuously exposed to low levels
of complement activation with deposition of the C5b-9 MAC but without
significant EC lysis.7-9 Although DAF, CD59, and MCP are
all expressed on ECs and contribute to the control of complement
activation on the cell surface,10-12 the specific contributions of these different molecules to the protection of the
endothelium during inflammation remain to be determined.2
Exposure of ECs to proinflammatory cytokines results in marked
phenotypic changes including upregulation of adhesion molecules, secretion of soluble mediators, and changes in vascular tone and permeability.13,14 In addition, it is well established that complement-activation products generated during an inflammatory response may act directly on ECs and influence their
function.15 Thus, C1q may upregulate the expression of
E-selectin and intercellular adhesion molecule-1
(ICAM-1),16 while C5a has been shown to induce P-selectin
expression by ECs.17 Furthermore, it has been reported that
sublytic concentrations of C5b-9 can bind to ECs, stimulating the
expression of adhesion molecules and the release of biologically active
mediators. Thus, assembly of the MAC on the EC surface may upregulate
the expression of tissue factor,18,19 as well as P- and
E-selectin, vascular cell adhesion molecule-1 (VCAM-1), and
ICAM-1.19-21 In addition, the secretion by ECs
of an array of soluble mediators in response to C5b-9, including the
chemokines interleukin-8 (IL-8) and monocyte-chemoattractant protein-1,22 prostacyclin,23 von Willebrand
factor,20 and the growth factors basic fibroblast growth
factor and platelet-derived growth factor,24 has been reported.
We aimed to investigate the hypothesis that the maintenance of vascular
integrity in the face of persistent complement activation during
chronic inflammation is dependent on the induction of one or more of
the complement-regulatory proteins DAF, CD59, and MCP by
proinflammatory mediators. Thus, soluble and cell-associated mediators
of the inflammatory response might act to upregulate the innate
protective mechanisms of endothelium against the potentially deleterious effects of complement activation, in parallel with their
ability to facilitate leukocyte adhesion and transmigration. Although
there is some evidence that DAF, MCP, and CD59 can be upregulated on
the EC surface in vitro, the mechanisms of control remain poorly
understood. The expression of DAF on human umbilical vein ECs (HUVECs)
may be upregulated by phorbol esters,25 wheat germ
agglutinin,26 and histamine.27 Despite this,
there are few data on the regulation of complement-regulatory proteins
by cytokines on ECs, with the exception of one report showing a modest increase in DAF expression by HUVECs in response to stimulation with
IL-4 or IL-1.28 To our knowledge, there is no published information on the regulation of the expression of
complement-regulatory proteins by microvascular ECs, which may be
particularly vulnerable to complement activation during inflammatory
responses that develop in the tissues in vivo. In this study, we
characterized the expression and regulation of DAF, MCP, and CD59 on
cultured human large- and small-vessel ECs. We provide evidence that
DAF is inducible on the EC surface, by both protein kinase C
(PKC)-dependent and -independent pathways, in response to soluble and
cell-associated proinflammatory mediators, and demonstrate a functional
role for this response in the protection of EC viability.
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MATERIALS AND METHODS |
Monoclonal antibodies and other reagents.
The monoclonal antibodies (MoAbs) included the following. A35 (IgG1)
against CD59 and anti-DAF MoAbs 5B2 and 1C6 (both IgG1); the latter
were kind gifts from Dr T. Fujita (University of Tsukuba, Tsukuba,
Japan). Anti-DAF MoAb 1H4 (IgG1) and TRA-2-10 (IgG1) against MCP were
kind gifts from Dr D.M. Lublin and Professor J.P. Atkinson,
respectively (Washington University School of Medicine, St Louis, MO).
The anti-endoglin (CD105) mAb RMAC8 (IgG2a) and the anti-class I MoAb
W6/32 (IgG2a) were kind gifts from Dr A. d'Apice (St Vincent's
Hospital, Victoria, Australia) and Dr N. Davey (Imperial College School
of Medicine, London, UK), respectively. IgG1 and IgG2a control MoAbs
were obtained from the adhesion panel of the Fifth CD Workshop, and the
EC-specific mAb EN4 (IgG1) was from Sanbio (Uden, Holland). Recombinant
human tumor necrosis factor- (TNF-) and the mAb MOPC-21, which
was used as an isotype-matched negative control, were kind gifts from
Dr Martyn Robinson (Celltech, Slough, UK). We are grateful for the gift
of the following cytokines: recombinant human interferon gamma
([IFN-] Glaxo, Geneva, Switzerland), IL-4 (Immunex Corp, Seattle,
WA), and IL-1 (Glaxo, Geneva, Switzerland). The specific PKC
antagonist RO31-822029 was a kind gift from Dr Trevor
Hallam (Roche, Welwyn Garden City, UK). The PKC agonist phorbol 12, 13-dibutyrate (PBu), bovine serum albumin (BSA), nonenzymatic cell-dissociation solution, and cycloheximide were purchased from Sigma
Chemical Co (Poole, UK). Normal human serum (NHS) was obtained from
blood samples from healthy individuals, collected under sterile conditions into glass tubes and allowed to clot at 37°C prior to
further incubation at 4°C for 2 hours to induce clot retraction. Following centrifugation at 1,800g, the NHS was collected,
pooled, and stored at  70°C.
Cell isolation and culture.
HUVECs were isolated from umbilical cords by digestion with collagenase
type II (Boehringer Mannheim, Lewes, UK) as described previously,30 and cultured in 1% gelatin-coated tissue
culture flasks (Costar, Cambridge, MA) in medium 199 ([M199] ICN
Biomedicals Inc, Costa Mesa, CA) supplemented with 20% fetal bovine
serum ([FBS] Hyclone Laboratories Inc, Logan, UT), 100 IU/mL
penicillin, 0.1 mg/mL streptomycin, 2 mmol/L L-glutamine
(all from GIBCO-BRL Life Technologies, Paisley, UK), 10 U/mL heparin
(Leo Laboratories, Prince Risborough, UK), and 30 µg/mL EC growth
supplement (Sigma). Dermal microvascular ECs (DMECs) were isolated from
human foreskins and cultured in fibronectin-coated flasks as previously
described in detail.31 Each experiment was performed with
ECs at passage 3 to 6. The human dermal microvascular cell line
HMEC-1,32 a kind gift from Dr E. Ades (Centers for Disease
Control, Atlanta, GA), was cultured in 1% gelatin-coated tissue
culture flasks in MCDB-131 growth medium (GIBCO-BRL) supplemented with
10% FBS, 100 IU/mL penicillin, 0.1 mg/mL streptomycin, 2 mol/L
L-glutamine, and 10 ng/mL epidermal growth factor (Becton
Dickinson, Bedford, MA).
Treatment of cells with phosphatidylinositol-specific phospholipase
C.
Phosphatidylinositol-specific phospholipase C (PIPLC) was prepared from
Bacillus cereus by Dr Peter Robinson (Imperial College School
of Medicine, London, UK). To cleave the glycosyl-phosphatidylinositol (GPI)-anchored proteins DAF and CD59 from the cell surface, monolayers of ECs in 35-mm petri dishes (6 × 105 cells per dish)
were washed three times with Hanks' balanced salt solution (HBSS)
before addition of PIPLC diluted 1:100 in M199. After incubation at
37°C for 30 minutes, ECs were washed with HBSS, harvested with
trypsin/EDTA (ICN Biomedicals), and analyzed by flow cytometry for
surface antigen expression.
Flow cytometry.
Following stimulation with test factors, monolayers of ECs were
harvested by exposure to trypsin/EDTA for 1 minute at 37°C. After
repeated pipetting to ensure single-cell suspensions, the cells were
stained with the appropriate primary MoAb for 30 minutes at 4°C.
After washing twice in phosphate-buffered saline/2.5% FBS, ECs were
resuspended in fluorescein isothiocyanate (FITC)-labeled rabbit
anti-mouse Ig (DAKO, Glostrup, Denmark) for 30 minutes at 4°C,
followed by washing as before and fixation in 1% paraformaldehyde. Samples were analyzed on an Epics XL-MCL flow cytometer (Coulter, Hialeah, FL) by counting 10,000 cells per sample. In some experiments, results are expressed as the relative fluorescence intensity (RFI), which represents the mean fluorescence intensity (MFI) with test MoAb
divided by the MFI using an isotype-matched irrelevant MoAb. In the
inhibition experiments, RO31-8220 and cycloheximide led to a small
reduction in the constitutive expression of DAF by resting ECs. To
control for this effect, the results of these experiments are expressed
as the RFI ratio, calculated as follows: RFI ratio = DAF RFI on EC
Treated With Proinflammatory Stimulus/DAF RFI on Unstimulated EC Control.
Northern blotting analysis.
Confluent ECs in 75-cm2 tissue culture flasks were
incubated with TNF- (10 ng/mL) and IFN- (500 U/mL) or plain
medium alone for a maximal 24 hours at 37°C. At the end of the
stimulation, cells were lysed in guanidinium isothiocyanate (Sigma) and
RNA was extracted as described by Chomczynski and Sacchi.33
Purified RNA was resuspended in 20 µL RNase-free water and stored at
70°C before use. The probe for DAF was obtained from a plasmid
vector34 and the DAF insert released by incubation for 2 hours at 37°C with the SalI and XbaI restriction
enzymes. Northern blotting was performed as described by Sambrook et
al.35 Purified RNA was run on 1% formaldehyde agarose
gels, blotted onto GeneScreen membrane (DuPont, Letchworth, UK), and
fixed in a UV Crosslinker (Stratagene, Cambridge, UK). Membranes were
prehybridized for 4 hours in 1 mol/L NaCl with 50% (vol/vol)
formamide, 4 mg DNA (salmon tests), 0.1% (wt/vol) sodium dodecyl
sulfate (SDS), and 10% (wt/vol) dextran sulfate (all from Sigma).
Approximately 200 ng purified probes were boiled for 3 minutes and used
for radiolabeling with 32P-dCTP using the Klenow fragment
of Escherichia coli DNA polymerase (Northumbria Biologicals,
Cramlington, UK). Membranes were hybridized to appropriate
32P-dCTP-labeled cDNA probes overnight at 42°C, and were
then washed in solutions of 0.1% SDS (wt/vol) containing successively
lower concentrations of SSC buffer (1× SSC is 0.15 mol/L NaCl plus
0.015 mol/L sodium citrate, pH 7). Specific hybridization was detected by autoradiography following exposure to Kodak XOMat film (Eastman Kodak, Rochester, NY). For quantification, Northern blots
were scanned using the Appligene Image Analysis System (Appligene, Durham, UK), and densitometry was performed using National Institutes of Health Image 1.52 software (Bethesda, MD). Values were
corrected for ethidium bromide-stained rRNA-loading patterns, and an
arbitrary value of 1 was assigned to unstimulated ECs.
Generation of the MAC.
ECs were plated at confluence (2 × 105 cells per well)
in 24-well plates (Costar) and cultured overnight at 37°C. They were then opsonized with MoAb RMAC8 (IgG2a), negative control MoAb, or plain
medium alone for 1 hour prior to addition of 2.5% NHS, heat-inactivated human serum ([HIHS] 30 minutes at 56°C), or C7- or
C8-deficient serum (Sigma) followed by incubation at 37°C for up to
48 hours. At the end of the assay, the EC monolayers were washed and the cells were harvested by trypsin/EDTA. After repeated pipetting to ensure single-cell suspensions, the cells were stained with an FITC-labeled anti-DAF MoAb (Serotec, Kidlington, Oxford, UK)
for 30 minutes at 4°C, and DAF expression was then analyzed by flow
cytometry as described before. The serum concentrations in
the assay were defined as sublytic using a standard
51Cr-release assay12 (data not shown). In
addition, in all experiments, cell viability was assessed by
examination of cell monolayers using phase-contrast microscopy,
cell-counting, and estimation of trypan blue exclusion in the EC
populations before staining.
C3-binding assay.
ECs were plated in 35-mm petri dishes (6 × 105 cells
per dish) and cultured overnight at 37°C before addition of TNF
(10 ng/mL) and IFN (500 U/mL), or in plain medium alone for 48 hours. Following harvesting with trypsin/EDTA, ECs were pelleted in
96-well v-bottom plates (Costar) and incubated with RMAC8 MoAb for 30 minutes at 4°C. After washing with HBSS/1% BSA, ECs were incubated
with 100 µL 20% NHS in M199 for 2 hours at 37°C prior to washing
with HBSS/1% BSA and addition of FITC-conjugated rabbit anti-human
C3c (DAKO) at 1:40 dilution for 30 minutes at 4°C. After washing
twice with HBSS/1% BSA, the presence of C3c was estimated using
flow-cytometric analysis as already described. Control samples included
the omission of NHS and the addition of 10 mmol/L EDTA to the NHS to
inhibit the classic pathway of complement activation. In the inhibition studies, the blocking MoAbs were added to the assay with the RMAC8 MoAb
to achieve a final concentration of 50 µg/mL.
Statistics.
Differences between the results of experimental treatments were
evaluated by the Mann-Whitney U test. Differences were
considered significant at a P value of <.05.
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RESULTS |
Expression of DAF, MCP, and CD59 on ECs.
Flow-cytometric analysis of resting ECs demonstrated that DAF, MCP, and
CD59 are constitutively expressed on the cell surface of both HUVECs
and DMECs. All 3 molecules were found on greater than 95% of resting
ECs with unimodal expression (Fig 1A). On large-vessel ECs, CD59 and
DAF are known to be attached to the cell surface via a GPI
anchor,2 which may influence the function of these
molecules.3 To investigate the membrane-anchoring of DAF
and CD59 on microvascular ECs, the expression of both molecules was
compared before and after treatment of ECs with PIPLC. Incubation of
ECs with PIPLC for 30 minutes at 37°C removed greater than 90% of
DAF and CD59 from the surface of HUVECs and DMECs, demonstrating that
these molecules are predominantly GPI-anchored on both cell types (Fig
1B). In contrast, MCP, although constitutively expressed on both HUVECs
and DMECs, was not released by PIPLC, consistent with attachment to the
cell surface via a transmembrane anchor (data not shown).4
The specificity of PIPLC for GPI-anchored proteins alone was further
confirmed by its failure to alter the surface expression of CD31 and
ICAM-1 (data not shown). To exclude a deleterious effect of trypsin on
the levels of DAF, CD59, and MCP measured, HUVECs were harvested with
either trypsin/EDTA or a nonenzymatic cell-dissociation solution
(Sigma) and analyzed by flow cytometry. No difference in the expression
of DAF, CD59, or MCP was observed following the different treatments
(Fig 1C).
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| Fig 1.
Expression of DAF, CD59, and MCP on HUVECs and DMECs. (A)
Expression of DAF, CD59, and MCP on resting ECs was assessed by flow
cytometry using mAbs 5B2, A35, and TRA-2-10, respectively. The figure
shows expression on DMECs and HUVECs with background fluorescence
(FITC-labeled rabbit anti-mouse Ab alone, ...) and specific
antigen expression ( ). (B) EC monolayers were incubated for 30 minutes in the absence or presence of PIPLC before harvesting and
analysis by flow cytometry. DAF and CD59 were detected with MoAb 5B2
and A35, respectively. The figure shows background fluorescence
(...), untreated ECs demonstrating constitutive expression (shaded
histogram), and PIPLC-treated ECs (). (C) HUVECs were harvested with
either trypsin/EDTA ( ) or a nonenzymatic cell-dissociation solution
( ) before analysis by flow cytometry. DAF, CD59, and MCP were
detected with MoAbs 5B2, A35, and TRA-2-10, respectively. The figure is
representative of 3 similar experiments.
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DAF is cytokine-inducible on ECs.
To investigate the regulation of complement-regulatory proteins on the
EC surface during inflammation, monolayers of ECs were stimulated with
the proinflammatory cytokines TNF-, IL-1, IFN-, or a
combination of TNF- and IFN- and subsequently analyzed by flow
cytometry. The concentrations of cytokines were based on those
previously demonstrated to induce EC activation and upregulation of
surface molecules including E-selectin, VCAM-1, ICAM-1, and Thy-1,30,31,36 and dose-response curves for each cytokine tested were generated (data not shown). No significant increase in the
expression of either CD59 (Fig 2C and D) or MCP (Fig 2E and F) was
found following a 48-hour stimulation with these cytokines. Furthermore, an extended time course up to 72 hours showed similar results. However, a significant increase in DAF was found following stimulation for 48 hours with TNF- (DMECs and HUVECs) and IFN- (DMECs) (P < .05; Fig 2A and B). Moreover, the combination
of TNF- and IFN- was more effective than either cytokine alone, inducing an increase in DAF expression of up to 4-fold
(P < .02; Fig 2A and B). While we were able to consistently
demonstrate a small increase in DAF expression on HUVECs after 72 hours
of stimulation with IL-4, this did not reach significance. Furthermore, IL-4 had no effect on the expression of DAF on DMECs in 4 separate experiments on different EC preparations (data not shown). In addition,
we were unable to demonstrate an effect of IL-4 on the TNF-induced
expression of DAF or an effect of IL-4 on MCP or
CD59.
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| Fig 2.
Analysis of DAF, CD59, and MCP expression on DMECs and
HUVECs following stimulation with cytokines. Monolayers of ECs were
incubated for 48 hours in the presence or absence of IL-1 (10 ng/mL), IFN (500 U/mL), TNF- (10 ng/mL), or TNF- and IFN-
before harvesting and analysis by flow cytometry. DAF, CD59, and MCP
were detected using MoAbs 5B2, A35, and TRA-2-10, respectively. The
results (mean ± SD) are expressed as the RFI (MFI of test sample
divided by MFI of irrelevant isotype-matched negative control) for (A,
B) DAF, (C, D) CD59, and (E, F) MCP. The results are representative of
4 experiments performed on different EC lines. *P < .05, **P < .02. The efficacy of cytokines used in each experiment
was confirmed by their ability to induce VCAM-1 or ICAM-1 as
appropriate (data not shown). US, unstimulated.
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The flow-cytometric histograms of DAF expression 48 hours
poststimulation with a combination of TNF- and IFN-, show an
increase in the MFI of up to 4-fold for both HUVECs and DMECs (Fig
3A). To investigate the kinetics of this
response, flow cytometry was performed on EC monolayers stimulated for
24, 48, and 72 hours with TNF- and IFN-. An increase in DAF was
first detectable after 24 hours of stimulation (Fig 3B), and was
maximal by 72 hours and persisted to 96 hours (data not shown).
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| Fig 3.
Time course of DAF upregulation on HUVECs and DMECs by
cytokines. EC monolayers were stimulated for up to 72 hours with plain
medium or a combination of TNF- (10 ng/mL) and IFN- (500 U/mL).
After harvesting, single-cell suspensions were analyzed by flow
cytometry for DAF expression using MoAb 5B2. (A) Upregulation of DAF on
DMECs and HUVECs after 48 hours of stimulation with a combination of
TNF- and IFN-: negative control (...), constitutive
expression of DAF (), and TNF- and INF--induced DAF (filled
histograms). (B) Kinetics for DAF expression on DMECs and HUVECs
following stimulation with TNF- and IFN-. The figure is
representative of 3 similar experiments on each cell type.
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It is known that the activation of PKC in HUVECs by phorbol esters
results in the upregulation of DAF on the cell surface.25 To investigate whether PKC activation is required for the induction of
DAF on cytokine-stimulated ECs, we used the specific PKC antagonist RO31-8220,29 which we have previously demonstrated to be an effective and nontoxic inhibitor of EC PKC.31,36 In these
experiments, the HMEC-1 cell line was used and the cells were
pretreated for 30 minutes with RO31-8220 before stimulation with
TNF- and IFN- or PBu. The induction of DAF following 48 hours of
stimulation with TNF and IFN- was not inhibited by RO31-8220,
suggesting cytokine signaling via a PKC-independent pathway (Fig 4A).
In contrast, the induction of DAF by PBu was completely inhibited by
the presence of RO31-8220 (Fig 4B). In some experiments, treatment with
RO31-8220 led to a small reduction in the constitutive expression of
DAF, and the results are thus expressed as the RFI ratio to control for
this effect as detailed earlier. RO31-8220 did not alter the
constitutive expression of MCP, CD59, or
EN4.
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| Fig 4.
Effect of cycloheximide and the PKC antagonist
RO31-8220 on DAF expression by ECs. HMECs plated at confluence in
35-mm petri dishes (6 × 105 cells/dish) and
cultured overnight at 37°C were treated with (A, B) RO31-8220 (1 µmol/L) and (C) cycloheximide (CHX, 1 µg/mL) for 30 minutes before
addition of activating factors. The cells were subsequently stimulated
with (A, C) TNF- (10 ng/mL) and IFN- (500 U/mL), (B) PBu (50 ng/mL), or plain medium for 48 hours. After harvesting, DAF expression
was measured by flow cytometry using MoAb 5B2, with MOPC-21 as an
isotype-matched negative control. The figure shows the effect of (A, B)
specific PKC antagonist RO31-8220 on TNF- and IFN-- and
PBu-induced DAF expression, respectively, and (C) CHX on TNF-
and IFN--induced DAF. CHX also completely inhibited PBu-induced DAF
(data not shown). To exclude the effects of nonspecific cytotoxicity,
cell viability was assessed by monolayer morphology,
cell-counting, and trypan blue exclusion on EC populations before
staining (data not shown). To control for any effect of the antagonists
on the constitutive expression of DAF, results are expressed as the RFI
ratio. The figure is representative of 4 similar experiments.
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Cytokine-induced DAF on ECs requires protein synthesis.
In experiments similar to those already described, HMECs were
pretreated for 30 minutes with cycloheximide before stimulation with
TNF- and IFN- or PBu. The presence of cycloheximide completely inhibited the cytokine-induced (Fig 4C) and PBu-induced (data not
shown) expression of DAF, suggesting that the induction of DAF is
dependent on de novo protein synthesis. Northern blot analysis was
performed using mRNA extracted from unstimulated ECs and cells stimulated with TNF- and IFN- for up to 24 hours. DAF mRNA was detectable at a low level in unstimulated ECs and was clearly upregulated following a 6-hour stimulation with cytokines in both DMECs
and HUVECs (Fig 5). Two DAF mRNA transcripts were detected at 2.4 and
1.8 kb, as previously reported.34,37 Quantification of mRNA
levels in resting and cytokine-stimulated ECs using densitometric scanning demonstrated a 7-fold increase in DMECs and a 4-fold increase
in HUVECs following 6 hours of stimulation with TNF- and IFN-.
After cytokine stimulation, the observed increase in DAF mRNA was first
detectable at 3 hours and maximal at 6 to 8 hours, and declined to
basal levels by 24 hours poststimulation (data not shown). Thus, the
upregulation of cell-surface expression following stimulation of ECs
with cytokines is associated with a transient increase in DAF
steady-state mRNA.
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| Fig 5.
Northern blot analysis of cytokine-induced DAF on HUVECs
and DMECs. ECs were cultured for 6 hours in plain medium alone or
supplemented with a combination of TNF- (10 ng/mL) and IFN- (500 U/mL). Total RNA was isolated and Northern blots were prepared. Lane 1, unstimulated DMECs; lane 2, TNF- and IFN--stimulated DMECs; lane
3, unstimulated HUVECs; lane 4, TNF- and INF--stimulated HUVECs.
The ethidium bromide-stained gel confirmed equal loading of RNA in
each lane. These findings were confirmed in 3 separate experiments.
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Assembly of the MAC on ECs induces DAF expression.
Endothelial cells at sites of inflammation may be activated by the MAC
of complement and by proinflammatory cytokines. The effect of the MAC
on EC DAF expression was studied by generating the C5b-9 complex at
levels that were shown in our preliminary experiments to be sublytic to
ECs. The EC monolayers were opsonized with an IgG2a anti-endoglin
(CD105) MoAb RMAC8.38 IgG2a is the optimal murine isotype
for C fixation, and endoglin was chosen as the target antigen because
it is highly expressed on the EC surface and its expression is not
upregulated by cytokines.39 Following
opsonization, ECs were incubated with 2.5% NHS at 37°C to induce generation of the MAC. Exposure of HMECs to the MAC increased
DAF expression on the cell surface up to 3-fold (Fig 6). Kinetic
analysis demonstrated that this response was maximal at 24 hours and
was maintained at 48 hours. Similar responses were observed at the same
time points with HUVECs (data not shown). The dependence on C
activation and the generation of the C5b-9 complex was demonstrated by
the lack of response when either HIHS or C7-deficient serum were used
instead of NHS (Fig 6). Furthermore, no response was found with
C8-deficient serum, suggesting that generation of the C5b-7 complex is
insufficient to induce DAF (data not shown). As with
cytokine-stimulated DAF expression, inclusion of the PKC antagonist
RO31-8220 did not inhibit MAC-induced DAF. It should be noted that the
RFIs in these experiments were lower than those described earlier,
since a directly FITC-labeled anti-DAF MoAb was used for the detection
of DAF.
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| Fig 6.
Upregulation of DAF on ECs following assembly of the MAC
of complement. HMECs were plated at confluence in 24-well plates
(2 × 105 cells/dish) and cultured overnight at 37°C.
They were then opsonized with MoAb RMAC8 or negative control MoAb for 1 hour, followed by addition of 2.5% NHS to induce formation of the MAC.
In parallel wells, ECs were pretreated with a combination of TNF-
and IFN- for 16 hours before assembly of the MAC. Additional
controls included the addition of HIHS or C7-deficient serum in place
of NHS. To inhibit PKC, RO31-8220 was added for 30 minutes before
assembly of the MAC and remained in the cultures throughout the assay.
After a total incubation of 24 hours at 37°C, ECs were harvested and
DAF expression was quantified by flow cytometry using an FITC-labeled
anti-DAF MoAb.
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To investigate the combined effects of the MAC and proinflammatory
cytokines on EC DAF expression, EC monolayers were treated with TNF-
and IFN- overnight prior to washing and opsonization as before.
Pretreatment of ECs with cytokines followed by MAC generation induced
an increase of DAF in excess of 3-fold, greater than the increase found
with either the MAC or cytokines alone (Fig 6). This effect was
dose-dependent (data not shown) and was maximal following treatment of
ECs with 10 ng/mL TNF- and 250 U/mL IFN-. Inclusion of
C7-depleted serum instead of NHS did not enhance the response, and the
increase in DAF levels was the same as observed following treatment
with the cytokines alone. As predicted by the results described,
inhibition of PKC by RO31-8220 did not abrogate this response (data not shown).
Cytokine-induced DAF reduces complement binding to ECs.
The binding of complement factor C3 to the EC surface was used to
investigate whether the observed induction of DAF could afford
increased protection to ECs against complement-mediated injury. HMEC
monolayers were opsonized with RMAC8 followed by incubation with 20%
NHS for 2 hours, after which C3 binding to the cell surface was
quantified by flow cytometry using a FITC-labeled antibody against C3c.
The addition of 10 mmol/L EDTA to the NHS completely inhibited C3
binding, confirming that the response was dependent on the classic
pathway of complement activation (data not shown). Moreover,
stimulation of ECs for 48 hours with TNF- and IFN- before
analysis reduced the deposition of C3 on the cell surface by 42%
compared with unstimulated cells (P = .014; Fig 7A). Parallel
flow-cytometric analysis confirmed that EC expression of endoglin was
not altered by cytokine stimulation (data not shown). To confirm the
role of DAF in the reduction of C3 binding following cytokine
stimulation, the inhibitory anti-DAF MoAb 1H4, which does not fix
complement (J.C.M. and D.O.H., unpublished observation, December 1998),
was included in the assay. In addition, we also studied
the effects of the non-complement-fixing, inhibitory anti-CD59 MoAb
A35,40 which would not be expected to inhibit C3 binding.
The addition of MoAb 1H4 markedly increased the binding of C3 to
unstimulated opsonized ECs exposed to 20% NHS (Fig 7B). Furthermore,
the reduction of C3 binding after 48 hours of prestimulation with
TNF- and IFN- was reversed by the presence of 1H4, with the
resultant detected level of C3 significantly greater than that found on
unstimulated ECs (P < .05). No significant increase in the
binding of C3 was detected in the presence of either anti-CD59 or
irrelevant control MoAbs. The inclusion of blocking MoAbs with DAF or
CD59 did not significantly increase cell lysis at this concentration of
NHS (data not shown). Therefore, these observations suggest that the
increased levels of cell-surface DAF observed following stimulation
with cytokines or the MAC may provide additional protection to ECs
against complement-mediated injury during
inflammation.
View larger version (33K):
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| Fig 7.
Functional analysis of cytokine-induced DAF on ECs. HMECs
were plated at confluence in 35-mm petri dishes
(6 × 105 cells/dish) and cultured overnight at 37°C.
They were then stimulated with a combination of TNF- (10 ng/mL) and
IFN- (500 U/mL) for 48 hours. After harvesting, ECs were incubated
with the anti-endoglin MoAb RMAC8 or plain medium alone for 30 minutes
at 4°C. In the blocking experiments, the anti-DAF MoAb 1H4, anti-CD59
MoAb A35, or isotype-matched negative control MoAb (final
concentration, 50 µg/mL) were added to the assay with RMAC8. The ECs
were then washed in HBSS/1% BSA before addition of 20% NHS for 2 hours at 37°C. Binding of C3 was detected by flow cytometry using
FITC-conjugated rabbit anti-human C3. (A) Percent C3 binding
(mean ± SD) to unstimulated () and TNF- and
IFN--stimulated ( ), HMECs, with binding to unstimulated ECs
shown as 100%: (B) changes in C3 binding (RFI mean ± SD) on
unstimulated () and TNF- and IFN--stimulated () HMECs in
the presence of non-complement-fixing inhibitory MoAbs 1H4 (anti-DAF)
and A35 (anti-CD59). *P < .05.
|
|
| |
DISCUSSION |
DAF is widely expressed on the surface of both hematopoietic and
nonhematopoietic cells, suggesting that it plays an important role in
protection against the toxic effects of complement
activation.3,41 The evidence to date suggests that the
surface expression of DAF may vary between tissues and may be regulated
by both physiologic and pathophysiologic
mechanisms.2,8,42,43 However, the understanding of the
regulation of DAF and its functional significance remains
incomplete.2 Our study demonstrates that exposure of ECs to
soluble cytokines and cell-associated MAC, which may coexist at sites
of inflammation, increases cell-surface expression of DAF. The data
suggest that these factors, which promote leukocyte adhesion to and
transmigration across the endothelium during inflammation, may also
protect ECs against complement-mediated injury in chronic inflammatory disease.
In the present study, initial experiments confirmed the presence of
DAF, MCP, and CD59 on HUVECs under basal culture
conditions.10,12 In addition, we found that all three
molecules are expressed at equivalent antigen densities on DMECs. The
constitutive expression of complement-regulatory proteins is thought to
protect the endothelium against the constant low-level activity of the
alternative pathway.3,12 However, there is a greatly
increased potential for local endothelial injury following activation
of the classic or alternative pathways of complement. Under these
circumstances, induction of complement-regulatory protein gene
expression might be expected to maintain or enhance the protection of
host tissues against complement-mediated injury. This hypothesis is
supported by our current observations that TNF- and IFN- induced
DAF expression on ECs. This upregulation was dependent on increased
steady-state mRNA and protein synthesis and was maximal at cytokine
concentrations previously shown to optimally induce ICAM-1 and VCAM-1
on ECs.30 The relatively delayed time course of DAF protein
upregulation on ECs is not unique and is comparable to that previously
reported for the Thy-1 and major histocompatibility complex class I and
II molecules on ECs.13,31 In contrast, we detected no
change in the basal level of either MCP or CD59 following EC treatment
with a variety of cytokines. Previous studies have demonstrated
upregulation of DAF expression on HUVECs in response to stimulation
with phorbol esters, wheat germ agglutinin, and
histamine.25-27 Moreover, Moutabarrik et al28
found a slight increase in DAF following stimulation of HUVECs with
IL-1, lipopolysaccharide, and IL-4, but not with TNF, and a small
increase in CD59 on TNF-stimulated cells. Although we also found a
small increase in DAF on HUVECs in response to treatment with IL-4,
this did not reach statistical significance and IL-4 had no effect on
DAF expression by DMECs in our studies.
In addition to cytokines, there is compelling evidence that
complement-activation products mediate changes in EC surface-antigen expression.15-17 Thus, the MAC is known to increase
TNF-induced expression of E-selectin and ICAM-1 on ECs and thereby to
increase neutrophil adhesion.19,22 This raised the question
of whether the MAC itself is capable of influencing the expression of
DAF on ECs. Using an in vitro model in which an anti-endoglin MoAb and
NHS were used to localize complement activation to the EC surface, we
observed a 3-fold increase in DAF, which was maximal 24 hours
postactivation. The dependence of this response on generation of the
C5b-9 complex was demonstrated by the failure of HIHS and C7- and
C8-deficient serum to induce DAF. The lack of response with
C8-deficient serum suggested that the complete C5b-9 complex is
required and that the previously described signaling capacity of the
C5b-7 complex44 is insufficient to induce DAF expression. While the response was consistently enhanced by prestimulation of ECs
for 16 hours with TNF- and IFN-, the effect of prestimulation with cytokines and the MAC together was typically additive rather than
synergistic, as previously described for ICAM-1 and
E-selectin.22
Our data therefore suggest that following activation of complement,
generation of C5b-9 can induce a feedback loop of vascular protection
via upregulation of DAF on the EC surface, a mechanism that is enhanced
by the presence of proinflammatory cytokines. This may be important in
human inflammatory diseases such as atherosclerosis, systemic lupus
erythematosus, rheumatoid arthritis, and glomerulonephritis, in which
complement-activation products may be deposited on the cell
surface.7-9,45,46 Furthermore, the capacity of the MAC to stimulate DAF expression during EC exposure to sublytic
concentrations of antibody and complement may contribute significantly
to the phenomenon of accommodation, in which the endothelium becomes resistant to complement-fixing alloreactive or xenoreactive antibodies in the transplantation setting.47
The role of PKC in the regulation of DAF was studied using the
PKC-specific inhibitor RO31-8220,29 which completely
abrogated the PBu-induced upregulation of EC DAF.25
However, the presence of RO31-8220 had no effect on the induction of
DAF by TNF- and IFN-. Furthermore, although PKC has been
implicated as one of a number of potential signaling pathways activated
by C5b-9,44,48,49 RO31-8220 had no effect on DAF induction
by the MAC. These and other data36 suggest the presence in
ECs of agonist-specific pathways for the regulation of surface
proteins, which in turn may have important functional implications. It
is therefore of particular interest that a recent study has shown that
activation of PKC in HUVECs not only upregulates cell-surface DAF but
also results in deposition of the molecule in the extracellular
matrix.50 We have made similar observations with DAF and
Thy-1 both in vitro (J.C.M. and D.O.H., unpublished observation,
January 1995) and in situ in human skin, where both molecules were
found in close association with the dermal vascular basement
membrane.31,51 However, treatment of HUVECs
with cytokines did not induce deposition of DAF in the extracellular
matrix.50 Together, these observations indicate the
presence of separate pathways for DAF induction in ECs, each of which
may be capable of mediating a specific distribution of the molecule
within the tissues.
The functional role of DAF on ECs was assessed by measurement of
surface-bound C3. Stimulation with TNF- and IFN- reduced C3
binding to the EC surface following activation of complement. Moreover,
inclusion of an inhibitory anti-DAF MoAb (1H4) in the assay showed that
this effect was mediated, at least in part, by increased cell-surface
expression of DAF. This suggests that the upregulation of DAF to the
levels observed after EC exposure to proinflammatory cytokines and/or
the MAC provides enhanced endothelial protection. DAF-mediated
cytoprotection may combine with other inducible protective mechanisms
likely to be active during inflammation, including an increase in the
synthesis of factors H and I, which specifically regulate alternative
pathway activation.52,53 These mechanisms may be of
particular importance in the light of recent data demonstrating that
treatment of ECs with cytokines may result in complement activation and
C3 deposition on the exposed subendothelial basement
membrane.54
The evidence to date suggests that the relative hierarchy in terms of
the physiologic importance of the complement-regulatory proteins is
CD59 > DAF > MCP.2 However, our observations suggest that DAF is particularly important in providing additional EC protection during subacute and chronic inflammation, and this may
reflect the inefficiency of terminal complement-component activation
and its absolute dependence on excess C3 activation.55 In
addition, the demonstration of distinct pathways for regulation raises
the possibility that DAF may have functions in EC biology above and
beyond decay-accelerating activity. DAF expression on the apical
surface of ECs in response to cytokines would place DAF in a position
to interact with leukocytes during their adhesion and transmigration to
inflammatory sites. Indeed, a recent study has demonstrated an adhesive
reaction between erythrocyte DAF and its ligand CD97, a molecule
expressed on activated T lymphocytes and monocytes.56
Finally, in light of the role of PKC in EC proliferation,57
it is possible that the PKC-dependent pathway of DAF expression may be
implicated in EC proliferation and angiogenesis.
In summary, we have demonstrated that DAF is expressed by cultured
large- and small-vessel ECs and is inducible on these cells by
PKC-dependent and -independent pathways. The increased level of DAF
observed after exposure of ECs to cytokines and the MAC increases the
level of protection against complement-mediated damage via a feedback
loop, suggesting that this pathway may play an important role in
maintaining vascular integrity during subacute and chronic inflammatory responses.
| |
ACKNOWLEDGMENT |
We are grateful to P. Kiely, P. Singh, L. Lovat, and M. McNamara for
collection of the foreskins and to the staff of the maternity unit of
Hammersmith Hospital for provision of the umbilical cords. We would
like to thank Mark Walport, Bernie Morley, Marina Botto, Tony d'Apice,
Teizo Fujita, Doug Lublin, and John Atkinson for their help in this study.
| |
FOOTNOTES |
Submitted January 21, 1999; accepted May 4, 1999.
Supported by the Arthritis Research Campaign, The Medical Research
Council (UK), and in part by a discretionary professorial award from
the BHF.
The publication costs of this
article were defrayed in part by
page charge payment. This article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. section
1734 solely to indicate this fact.
Address reprint requests to Justin C. Mason, PhD, The BHF
Cardiovascular Medicine Unit, National Heart and Lung Institute,
Imperial College School of Technology and Medicine, Hammersmith
Hospital, Du Cane Road, London W12 ONN, UK; e-mail: jmason{at}rpms.ac.uk.
| |
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1598 - 1607.
[Abstract]
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A. R. Kinderlerer, F. Ali, M. Johns, E. A. Lidington, V. Leung, J. J. Boyle, S. S. Hamdulay, P. C. Evans, D. O. Haskard, and J. C. Mason
KLF2-dependent, Shear Stress-induced Expression of CD59: A NOVEL CYTOPROTECTIVE MECHANISM AGAINST COMPLEMENT-MEDIATED INJURY IN THE VASCULATURE
J. Biol. Chem.,
May 23, 2008;
283(21):
14636 - 14644.
[Abstract]
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S. Shilo, S. Roy, S. Khanna, and C. K. Sen
Evidence for the Involvement of miRNA in Redox Regulated Angiogenic Response of Human Microvascular Endothelial Cells
Arterioscler Thromb Vasc Biol,
March 1, 2008;
28(3):
471 - 477.
[Abstract]
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R. Steinberg, O. A. Harari, E. A. Lidington, J. J. Boyle, M. Nohadani, A. M. Samarel, M. Ohba, D. O. Haskard, and J. C. Mason
A Protein Kinase C{epsilon}-Anti-apoptotic Kinase Signaling Complex Protects Human Vascular Endothelial Cells against Apoptosis through Induction of Bcl-2
J. Biol. Chem.,
November 2, 2007;
282(44):
32288 - 32297.
[Abstract]
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E. L. Campbell, N. A. Louis, S. E. Tomassetti, G. O. Canny, M. Arita, C. N. Serhan, and S. P. Colgan
Resolvin E1 promotes mucosal surface clearance of neutrophils: a new paradigm for inflammatory resolution
FASEB J,
October 1, 2007;
21(12):
3162 - 3170.
[Abstract]
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M. I. Leite, M. Jones, P. Strobel, A. Marx, R. Gold, E. Niks, J. J.G.M. Verschuuren, S. Berrih-Aknin, F. Scaravilli, A. Canelhas, et al.
Myasthenia Gravis Thymus: Complement Vulnerability of Epithelial and Myoid Cells, Complement Attack on Them, and Correlations with Autoantibody Status
Am. J. Pathol.,
September 1, 2007;
171(3):
893 - 905.
[Abstract]
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A. J. Hwa, R. C. Fry, A. Sivaraman, P. T. So, L. D. Samson, D. B. Stolz, and L. G. Griffith
Rat liver sinusoidal endothelial cells survive without exogenous VEGF in 3D perfused co-cultures with hepatocytes
FASEB J,
August 1, 2007;
21(10):
2564 - 2579.
[Abstract]
[Full Text]
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J. Wehner, C. N. Morrell, T. Reynolds, E. R. Rodriguez, and W. M. Baldwin III
Antibody and Complement in Transplant Vasculopathy
Circ. Res.,
February 2, 2007;
100(2):
191 - 203.
[Abstract]
[Full Text]
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E. A. Lidington, R. Steinberg, A. R. Kinderlerer, R. C. Landis, M. Ohba, A. Samarel, D. O. Haskard, and J. C. Mason
A role for proteinase-activated receptor 2 and PKC-{epsilon} in thrombin-mediated induction of decay-accelerating factor on human endothelial cells
Am J Physiol Cell Physiol,
December 1, 2005;
289(6):
C1437 - C1447.
[Abstract]
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K. Wenzel, J. Zabojszcza, M. Carl, S. Taubert, A. Lass, C. L. Harris, M. Ho, H. Schulz, O. Hummel, N. Hubner, et al.
Increased Susceptibility to Complement Attack due to Down-Regulation of Decay-Accelerating Factor/CD55 in Dysferlin-Deficient Muscular Dystrophy
J. Immunol.,
November 1, 2005;
175(9):
6219 - 6225.
[Abstract]
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N. A. Louis, K. E. Hamilton, T. Kong, and S. P. Colgan
HIF-dependent induction of apical CD55 coordinates epithelial clearance of neutrophils
FASEB J,
June 1, 2005;
19(8):
950 - 959.
[Abstract]
[Full Text]
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J. van Beek, M. van Meurs, B. A. 't Hart, H. P. M. Brok, J. W. Neal, A. Chatagner, C. L. Harris, N. Omidvar, B. P. Morgan, J. D. Laman, et al.
Decay-Accelerating Factor (CD55) Is Expressed by Neurons in Response to Chronic but Not Acute Autoimmune Central Nervous System Inflammation Associated with Complement Activation
J. Immunol.,
February 15, 2005;
174(4):
2353 - 2365.
[Abstract]
[Full Text]
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J. C. Mason, R. Steinberg, E. A. Lidington, A. R. Kinderlerer, M. Ohba, and D. O. Haskard
Decay-accelerating Factor Induction on Vascular Endothelium by Vascular Endothelial Growth Factor (VEGF) Is Mediated via a VEGF Receptor-2 (VEGF-R2)- and Protein Kinase C-{alpha}/{epsilon} (PKC{alpha}/{epsilon})-dependent Cytoprotective Signaling Pathway and Is Inhibited by Cyclosporin A
J. Biol. Chem.,
October 1, 2004;
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41611 - 41618.
[Abstract]
[Full Text]
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K. Yamada, T. Miwa, J. Liu, M. Nangaku, and W.-C. Song
Critical Protection from Renal Ischemia Reperfusion Injury by CD55 and CD59
J. Immunol.,
March 15, 2004;
172(6):
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[Abstract]
[Full Text]
[PDF]
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S.-H. Li, P. E. Szmitko, R. D. Weisel, C.-H. Wang, P. W.M. Fedak, R.-K. Li, D. A.G. Mickle, and S. Verma
C-Reactive Protein Upregulates Complement-Inhibitory Factors in Endothelial Cells
Circulation,
February 24, 2004;
109(7):
833 - 836.
[Abstract]
[Full Text]
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B. Bussolati, A. Ahmed, H. Pemberton, R. C. Landis, F. Di Carlo, D. O. Haskard, and J. C. Mason
Bifunctional role for VEGF-induced heme oxygenase-1 in vivo: induction of angiogenesis and inhibition of leukocytic infiltration
Blood,
February 1, 2004;
103(3):
761 - 766.
[Abstract]
[Full Text]
[PDF]
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B Anlar
Infection and multiple sclerosis
J. Neurol. Neurosurg. Psychiatry,
May 1, 2003;
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[Full Text]
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J. C. Mason, Z. Ahmed, R. Mankoff, E. A. Lidington, S. Ahmad, V. Bhatia, A. Kinderlerer, A. M. Randi, and D. O. Haskard
Statin-Induced Expression of Decay-Accelerating Factor Protects Vascular Endothelium Against Complement-Mediated Injury
Circ. Res.,
October 18, 2002;
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[Abstract]
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S. L. Young, B. A. Lessey, M. A. Fritz, W. R. Meyer, M. J. Murray, P. L. Speckman, and B. J. Nowicki
In Vivo and in Vitro Evidence Suggest That HB-EGF Regulates Endometrial Expression of Human Decay-Accelerating Factor
J. Clin. Endocrinol. Metab.,
March 1, 2002;
87(3):
1368 - 1375.
[Abstract]
[Full Text]
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E. A. Lidington, D. O. Haskard, and J. C. Mason
Induction of decay-accelerating factor by thrombin through a protease-activated receptor 1 and protein kinase C-dependent pathway protects vascular endothelial cells from complement-mediated injury
Blood,
October 15, 2000;
96(8):
2784 - 2792.
[Abstract]
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S. Chen, T. Caragine, N.-K. V. Cheung, and S. Tomlinson
CD59 Expressed on a Tumor Cell Surface Modulates Decay-accelerating Factor Expression and Enhances Tumor Growth in a Rat Model of Human Neuroblastoma
Cancer Res.,
June 1, 2000;
60(11):
3013 - 3018.
[Abstract]
[Full Text]
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J. C. Mason, E. A. Lidington, S. R. Ahmad, and D. O. Haskard
bFGF and VEGF synergistically enhance endothelial cytoprotection via decay-accelerating factor induction
Am J Physiol Cell Physiol,
March 1, 2002;
282(3):
C578 - C587.
[Abstract]
[Full Text]
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TOP
ABSTRACT
INTRODUCTION