(Journal of Leukocyte Biology. 2002;71:433-444.)
© 2002
by Society for Leukocyte Biology
RAGE-mediated neutrophil dysfunction is evoked by advanced glycation end products (AGEs)
Kate S. Collison*,
Ranjit S. Parhar*,
Soad S. Saleh*,
Brian F. Meyer*,
Aaron A. Kwaasi*,
Muhammad M. Hammami*,
Ann Marie Schmidt
,
David M. Stern
and
Futwan A. Al-Mohanna*
* Biological & Medical Research, King Faisal Specialist Hospital and Research Centre, Riyadh, Saudi Arabia; and
Departments of Physiology and Surgery, Columbia University, College of Physicians and Surgeons, New York, New York
Correspondence: Futwan Al-Mohanna, Ph.D., Biological & Medical Research, MBC 03, King Faisal Specialist Hospital & Research Centre, P.O. Box 3354, Riyadh 11211, Saudi Arabia. E-mail: futwan{at}kfshrc.edu.sa
 |
ABSTRACT
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The accumulation of advanced glycation end products (AGEs) in the
tissue and serum of subjects with diabetes has been linked to the
pathogenesis of vascular complications. Because diabetes may be also
complicated by increased susceptibility to recurrent infection, we
investigated the effects of AGEs on human neutrophils, because their
burst of activity immediately upon engagement of pathogens or other
inflammatory triggers is critical to host response. We demonstrate the
presence of receptor for advanced glycation end products (RAGE) at the
message and protein levels. We also demonstrate that AGE albumin (but
not control albumin) binds with high affinity to human neutrophils
(Kd of 3.7±0.4 nM). The binding was blocked
almost completely by excess soluble RAGE, anti-RAGE antibodies, or
antibodies to CML-modified albumin. AGE albumin induced a
dose-dependent increase in intracellular-free calcium as well as actin
polymerization. Further, AGE albumin inhibited transendothelial
migration and Staphylococcus aureus-induced but not
fMLP-induced production of reactive oxygen metabolite. Moreover,
although AGE albumin enhanced neutrophil phagocytosis of S.
aureus, it inhibited bacterial killing. We conclude
that functional RAGE is present on the plasma membrane of human
neutrophils and is linked to Ca2+ and actin
polymerization, and engagement of RAGE impairs neutrophil
functions.
Key Words: calcium ROM transmigration phagocytosis signal
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INTRODUCTION
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Advanced glycation end products (AGEs) are a heterogeneous group
of nonenzymatically glycated and oxidized species, which occur in vivo
when proteins and lipids are exposed to aldoses or are subject to
oxidant stress [1
, 2
]. Initially, early
glycation products, Schiff bases, and Amadori products are reversibly
formed whenever plasma-glucose levels are elevated. A small proportion
of these products undergo further slow, irreversible chemical
rearrangements to form AGEs, which accumulate in the vasculature under
conditions that are accelerated during hyperglycemia and when protein
turnover is delayed [1
]. A number of cellular receptors
for AGEs have been identified, and the best characterized is receptor
for AGEs (RAGE), a member of the immunoglobulin superfamily of
receptors [3
]. AGEs have been shown to interact with
RAGE in vitro on cultured endothelium, resulting in increased monolayer
permeability, thrombogenicity, and proliferation [4
] and
may thus contribute to the loss of vascular homeostasis. Abnormal
levels of tissue and serum AGEs have been demonstrated in diabetes, a
disease where vascular homeostasis is clearly compromised
[5
]. A further complication associated with some
diabetic patients is the loss of the bodys immune defense, resulting
in delayed wound healing [6
] and an increased
susceptibility to infection [7
].
Neutrophils, the predominant circulating leukocytes, are the bodys
first line of cellular defense against invading microorganisms. It is
well recognized that neutrophil function is impaired in diabetic
patients [8
9
10
11
12
], however the exact mechanism(s) for
this impairment is not fully understood and is likely to be
multifactorial [13
].
In this paper, we investigate the direct interactions of AGEs
with human naïve neutrophils. We show that RAGE is present on
the neutrophil plasma membrane and that its engagement by AGEs results
in a rapid calcium-dependent activation of human neutrophils that is
associated with aberrant signal processing and altered neutrophil
responses. Although AGE-treated neutrophils exhibit an increased
phagocytic index, their ability to kill the ingested bacteria is
compromised. Our data suggest that RAGE engagement may protect ingested
bacteria from killing, which raises the possibility of a novel
mechanism through which recurrent infections may occur under conditions
of hyperglycemia.
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MATERIALS AND METHODS
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Preparation of AGE albumin, carboxymethylysine (CML) albumin, and
soluble RAGE
AGE albumin was prepared by incubating sterile human serum
albumin (HSA; Sigma Chemical Co., St. Louis, MO; A3782) or bovine serum
albumin (BSA; Fraction V; Sigma Chemical Co.; A3059) with glucose (0.5
M) for 6 weeks at 37°C, as described previously [14
,
15
]. AGE albumin was characterized based on its binding
to cultured endothelial cells and purified AGE-binding proteins as
described previously [16
] and based on its fluorescence
[1
]. Control, nonglycated albumin was prepared in an
identical manner, except that glucose was omitted from the incubation
buffer. Where indicated, AGE albumin was radiolabeled by
125I using the lactoperoxidase method [17
].
The labeled protein was isolated on a Sephadex G25 column, and its
final specific activity was determined. Fluorescein isothiocyanate
(FITC)-AGE albumin was prepared by incubating AGE albumin (25 mg) with
FITC (5 mg) in a volume of 1 ml phosphate-buffered saline (PBS; pH 7.4)
for 18 h at 4°C. Following extensive dialysis with PBS (pH 7.4)
to remove unbound FITC, FITC-AGE albumin was stored at -30°C until
use. At a protein concentration of 2 mg/ml, there was no detectable
lipopolysaccharide according to the Limulus amebocyte assay (Sigma
Chemical Co.). Control, nonglycated HSA or BSA was labeled with FITC
(FITC albumin) in a similar manner to AGE albumin. CML albumin was
prepared as described previously [18
]. Briefly, 2 mg/ml
BSA or HSA (Sigma Chemical Co.; A3059 and A3782, respectively) was
incubated at 37°C for 24 h with 0.75 mM glyoxylic acid and 0.3
mM NaCNBH3 in a buffer of 0.5 mM sodium phosphate, pH 7.4,
followed by extensive dialysis against PBS at 4°C. The proteins were
freeze-dried, reconstituted in PBS, and dialyzed against PBS before
storing at -20°C until use. In all experiments described in this
manuscript, human and bovine AGE albumin behaved similarly, and unless
otherwise stated, AGE albumin should therefore mean human AGE albumin.
Soluble RAGE was prepared using the baculovirus system
[19
], and monospecific rabbit anti-human RAGE
immunoglobulin G (IgG) was prepared and characterized as described
[14
].
Isolation of human neutrophils
Human peripheral blood neutrophils were prepared by dextran
sedimentation of heparinized whole blood obtained from healthy donors
as described previously [20
]. Contaminating red blood
cells (RBCs) were removed by hypotonic lysis with isotonic
NH4Cl. The remaining cells were resuspended in Krebs-HEPES
medium (pH 7.4) containing 120 mM NaCl, 1.3 mM CaCl2, 1.2
mM MgSO4, 4.8 mM KCl, 1.2 mM
KH2PO4, 25 mM HEPES
(N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic
acid), and 0.1% BSA; they were purified further through
neutrophil-isolation medium (Cardinal Associates, Santa Fe, NM). Final
purity and viability were between 98% and 99%, as indicated by flow
cytometry (FACScan; Becton Dickinson, San Jose, CA) and trypan blue-dye
exclusion tests.
Isolation of human endothelial cells
Human aortic endothelial cells (HAECs) were isolated from
the thoracic aorta of donor hearts (according to International Review
Board policies and procedure) as follows: A section of the
thoracic aorta (left over from cardiac-transplant surgery) was incised
longitudinally and placed flat in a petri dish with the intraluminal
surface exposed. The endothelium was gently scraped off using a sterile
scalpel blade and placed in HEPES-buffered medium 199 (Sigma Chemical
Co.), supplemented with penicillin (100 U/ml), streptomycin (100
µg/ml), L-glutamine (1 mM), bovine brain extract (25 µg/ml),
heparin (15 U/ml), endothelial cell growth factor (25 µg/ml; Sigma
Chemical Co.), and 20% fetal bovine serum. Cells were centrifuged and
resuspended in the same medium, plated onto sterile culture dishes,
coated with 0.5% gelatin, and cultured at 37°C and 5%
CO2. Endothelial cells were characterized by their
cobblestone morphology and positive immunostaining with antibodies to
von Willebrand factor (F3520; Sigma Chemical Co.) and with acetylated
low-density lipoprotein (DiI-Ac-LDL; Biogenesis, Bournmouth, U.K.).
Cells were used from passages 210 in all experiments at a split ratio
of 1:3. To confirm that endothelial cells were not activated during
isolation and culture, interleukin-1 (IL-1)
levels in the
conditioned medium were measured using enzyme-linked immunosorbent
assay (ELISA; R&D Systems, Minneapolis, MN.). IL-1
levels were
consistently found to be negligible (<4 pg/ml). Lipopolysaccharide
(LPS) levels were also tested routinely in culture media before and
after experimentation and were found to be undetectable.
Detection of RAGE by reverse transcriptase-polymerase chain
reaction (RT-PCR)
Quantitative extraction of total RNA from 5 x
106 neutrophils (>98% purity) or an equivalent number of
mononuclear cells was performed using TRI Reagent (Molecular Research
Centre, Inc., Cincinnati, OH), according to the manufacturers
instructions. RT-PCR was used for semiquantitative analysis of
transcript levels. cDNA was synthesized from the total RNA
representative of neutrophils (2.5x105) using avian
myoblastosis virus RT (Promega, Madison, WI), according to the
manufacturers protocol. Sense and antisense primers for human RAGE
were 5'-AGCGGCTGGAATGGAAACTGAA-3' (M91211; nucleotides 140162) and
5'-CTACAGGAGAAGGTGGGACGGG-3' (M1211; reverse complement of nucleotides
605627), respectively. The total cDNA of each sample was amplified by
PCR in a final volume of 50 µl containing 100 ng each primer, 100 mM
dNTPs, 50 mM KCl, 10 mM Tris-HCl (pH 8.3), 1.5 mM MgCl2,
and 1 unit Taq polymerase (Pharmacia, Uppsala, Sweden). Thirty-five
cycles of denaturing for 1 min at 94°C, annealing for 1 min at
55°C, and extension for 1 min at 72°C were used for amplification.
PCR products were electrophoresed on 2% agarose gels and stained with
ethidium bromide.
Western blotting of neutrophil preparations
For Western blotting of membrane-enriched neutrophil
preparations, human neutrophils were isolated, pelleted, and
resuspended in buffer containing 100 mM KCl, 3 mM NaCl, 3.5 mM
MgCl2, 10 mM HEPES, pH 7.3, and protease inhibitors
phenylmethylsulfonyl fluoride (PMSF; 1 mM), chymostatin (5 µg/ml),
antipain (5 µg/ml), pepstatin A (5 µg/ml), and leupeptin (10
µg/ml). Postnuclear supernatant (PNS) was prepared, layered over a 15
ml cushion of the above buffer containing 41% sucrose in a
polycarbonate ultracentrifugation tube, and centrifuged at 90,000
g at 4°C for 60 min. The plasma membrane-enriched fraction
was recovered after aspiration of the upper fraction containing the
cytosol. The purity of the isolated fractions was determined by
measuring specific alkaline phosphatase activity in the PNS, cytosol,
and membrane fractions, which were assayed subsequently for
alkaline-phosphatase activity. Equal amounts of protein (120, 280, 420
ng) from PNS, cytosol, and membrane-enriched fractions were run on
sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE)
gels and were used for Western blotting with anti-RAGE antibody.
Binding studies
Binding of 125I-AGE albumin to human neutrophils was
studied using suspensions of neutrophils (2.5x105/well) in
Krebs-HEPES buffer. Briefly, wells were incubated with Krebs-HEPES
buffer in the presence of various concentrations of
125I-AGE albumin alone (total binding) or with a 200-fold
excess of unlabeled AGE albumin (nonspecific binding). Where indicated,
neutrophils were preincubated with rabbit polyclonal antibody to RAGE
or nonimmune rabbit serum for 2 h at 4°C. Following a 3-h
incubation at 4°C, cells were washed rapidly four times in
Krebs-HEPES buffer, and cell-bound 125I-AGE albumin was
counted using a gamma counter (CliniGamma 1272; LKB-Wallac, Turku,
Finland). Experiments were performed in triplicate, and results were
analyzed using GraphPad Prism software (GraphPad Software, San Diego,
CA). For binding experiments performed with FITC-labeled AGE albumin,
FITC-AGE albumin (60 µg) was added to formaldehyde-fixed neutrophils
(1x105/100 µl) in the absence or presence of a 200-fold
excess of unlabeled AGE albumin for 15 min at 37°C in PBS (pH 7.4).
The cells were then washed three times to remove unbound AGE albumin
and resuspended in PBS, and the fluorescence associated with the cells
was analyzed using flow cytometry (Becton Dickinson) or confocal
laser-scanning microscopy (Leica-Kaki, Saudi Arabia). Identical
experiments were performed with FITC albumin in the presence and
absence of a 100-fold excess of unlabeled albumin. Unless otherwise
stated, soluble RAGE and anti-RAGE polyclonal antibodies were used at a
final concentration to give 100-fold excess over AGEs.
Internalization of FITC-labeled AGE albumin was followed by
confocal microscopy
Briefly, FITC-labeled AGE albumin (0.36 µM) was added to the
Krebs medium bathing the neutrophils, which were adherent to glass
coverslips. The appearance of intracellular fluorescence was followed
up with time. In similar experiments, the cells were also labeled with
the lysosomal label LysoTracker Red DND (Molecular Probes, Eugene, OR),
according to the manufacturers instructions.
Measurement of intracellular-free calcium
Neutrophils were loaded with Fura 2-AM as described previously
[21
]. The cells were washed and allowed to adhere to
glass coverslips for 15 min at room temperature. Coverslips with
adherent neutrophils were rinsed in Krebs-HEPES buffer and secured in a
custom-designed coverslip holder placed on a temperature-controlled
microscope stage (33°C), where calcium measurements were performed on
individual cells stimulated with various doses of AGE albumin using the
ionVision dual excitation system (ImproVision, Coventry, UK) as
described previously [22
].
Actin-polymerization measurements
Actin polymerization was measured using flow cytometry as
described previously [23
]. Briefly, human neutrophils
(107/ml) were incubated at 37°C for 10 min in a stirred,
temperature-controlled chamber. Samples (100 µl) were drawn before
and at different time intervals after addition of 0.36 µM AGE
albumin, and cells were fixed immediately in formaldehyde (3.7%
formaldehyde for 15 min at room temperature) and washed extensively in
PBS before use. Neutrophils were permeabilized using
lysophosphatidylcholine (4 µg/ml; Sigma Chemical Co.) for 5 min at
room temperature and stained for actin using fluorescein phalloidin
(0.33 µM for 1 h at room temperature; Molecular Probes). The
fluorescent intensity of washed cells was measured using flow
cytometry.
The extent of AGE albumin-induced actin polymerization was confirmed by
SDS-PAGE, and immunoblotting of the actin associated with the Triton
X-100-insoluble cytoskeleton. SDS-PAGE of actin was performed as
described previously [23
]. Separated protein bands were
transferred electrophoretically onto nitrocellulose membranes (Bio-Rad,
Hercules, CA) and blots washed twice with distilled water, preblocked
in 3% nonfat milk in PBS for 1 h at room temperature, and
incubated with antiactin antibodies at 4°C for 18 h. Following
extensive washing with PBS containing 0.02% Tween-20, membranes were
incubated with horseradish peroxidase-conjugated secondary antibodies
(Santa Cruz Biotechnology, Santa Cruz, CA) for 1 h at room
temperature. After further washing in the detergent buffer above, actin
bands were detected using enhanced chemiluminescence (ECL; Amersham,
Aylesbury, UK). Where indicated, neutrophils were pretreated with
bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetate
acetoxymethyl ester (BAPTA-AM; 10 µM, 30 min, at room temperature) to
chelate-intracellular calcium.
Measurement of transendothelial migration
Transwell chambers (6.5 mm diameter/3.0 µm pore size; Corning
Costar, Cambridge, MA) were used to evaluate the migration of human
neutrophils through endothelial cell monolayers. HAECs
(5x105/well) were seeded on the upper chamber of the
transwell and allowed to form monolayers. One-hundred percent
confluence was achieved at 1218 h post-seeding. This was confirmed by
bright-field microscopy. 51Cr-labeled neutrophils
pretreated with BSA or AGE albumin (15 min at 37°C) neutrophils
(2.5x105 cells) were added on top of the endothelial
monolayer in the upper transwell chamber. Krebs-HEPES medium (1 mL)
containing 1 µM chemotactic-peptide formyl-Met-Leu-Phe (fMLP) was
added to the lower transwell chamber, and both chambers were incubated
at 37°C, 5% CO2 for 2 h. At the end of the
incubation period, migrated neutrophils were collected from the lower
chamber, and radioactivity associated with the cells was measured using
a gamma counter (1272; Clinigamma, Turku, Finland).
Measurement of reactive oxygen metabolites (ROM) production
The production of ROM by human neutrophils was measured using
luminol-dependent chemiluminescence (LDCL) as described previously
[24
25
26
]. Briefly, neutrophils (1x106/ml)
were incubated with 5-amino-2,3-dihydrophthalazine-1,4-dione (luminol,
11 µM) at 37°C for 5 min prior to addition of stimulus, and the
resultant change in luminescence was displayed on a chart recorder.
Measurement of phagocytosis
Phagocytosis of fluorescein-labeled heat-killed
Staphylococcus aureus was performed as described previously
[20
]. Briefly, S. aureus (strain SA 133;
American Type Culture Collection, Manassas, VA; 37235) were heat-killed
at 100°C (15 min), washed twice in PBS (pH 7.2), and adjusted to
5 x 1011/ml. Fluorescence labeling was performed by
incubating the bacteria with FITC (50 mM) at 4°C for 72 h.
Unbound FITC was removed by extensive dialysis in PBS. The effect of
AGE albumin on neutrophil phagocytosis was determined after incubation
of neutrophils (106/ml) with labeled bacteria (100-fold
excess) in the presence of various doses of AGE albumin for 1 h at
37°C. The suspension was diluted tenfold and viewed under
fluorescence microscopy. The number of labeled bacteria ingested per
neutrophils was determined, and the results were expressed as change in
phagocytic index (PI), where PI was calculated by the following
formula: Number of bacteria in neutrophils counted in 50 fields of
view/number of neutrophils. The fluorescence of noningested bacteria
was quenched by the addition of 0.5% trypan blue to the cell
suspension before viewing.
Intracellular killing of ingested bacteria was measured as follows:
Neutrophils (105) were allowed to adhere on glass
coverslips for 15 min at 37°C. Unattached cells were removed by
washing with Krebs-HEPES medium. The cells were then treated with AGE
albumin or controlled albumin (1.1 µM) for 1 h at 37°C. Cells
were washed, and 1010 live S. aureus were added.
Phagocytosis was allowed to proceed for 1 h at 37°C. The number
of live bacteria was measured using a live/dead BacLight bacteria
viability kit (Molecular Probes) according to the manufacturers
instructions. To validate the live/dead bacteria assay, we performed a
series of experiments in which colony-forming ability of ingested
bacteria was investigated. Briefly, neutrophils were incubated with
live bacteria at a neutrophil:bacteria ratio of 1:100 for 1 h at
37°C in the presence of AGE albumin or albumin. To separate cells
from noningested bacteria, the samples were centrifuged through
neutrophil-isolation medium (NIM; lot #44706; Cardinal Associates) for
20 min at 1400 g and 4°C. The cells were collected,
washed, and resuspended in 1 ml Krebs-HEPES medium, and 25 µl samples
were spotted onto blood agar plates (lot #114260; Saudi Prepared Media
Lab., Riyadh, Saudia Arabia). The plates were incubated at 37°C for
72 h.
In some experiments, phagolysosome formation was measured by labeling
lysosomes with LysoTracker Red DND (50 nM for 15 min at RT) before
commencement of phagocytic assay, and cells were viewed under confocal
micrsocopy.
 |
RESULTS
|
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Is RAGE present in human naïve neutrophils
Because RAGE is the predominant receptor for AGE albumin in many
cell types, we tested the presence of RAGE mRNA by RT-PCR, which
revealed the presence of RAGE amplicons in human neutrophils
(Fig. 1 a
). Because the neutrophil preparations are typically 9899% pure
[as detected by fluorescein-activated cell sorter (FACS) analysis]
with 12% contamination of other cell types, namely peripheral blood
mononuclear cells (PBMC), the possibility existed that the RAGE
amplicons revealed by RT-PCR were a product of the contaminating PBMC
rather than neutrophils. We tested this by running RT-PCR of purified
neutrophils and various numbers of PBMC prepared from the same blood
donor. Under these conditions and even with 5 x 104
PBMC equivalent (representing 20% contamination), the amount of RAGE
amplicons was less than that obtained from the routinely prepared
neutrophil (unpublished results), suggesting that RAGE amplicons are
derived predominantly from mRNA of neutrophil origin. To test whether
this message is expressed, confocal microscopy using indirect
immunofluorescence of monospecific anti-RAGE IgG, which revealed
membrane-associated fluorescence that was not observed in control
experiments with isotypic IgG (Fig. 1b)
, was used. This was further
confirmed by Western blotting experiments in which RAGE was detected
predominantly in plasma membrane preparations isolated from neutrophil
homogenates (Fig. 1b)
.

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Figure 1. Presence of mRNA transcript in human neutrophils. (a) RT-PCR of RAGE
mRNA transcripts from human neutrophils. Molecular weight (MW) markers
in lane 1; negative control in lane 2; RAGE mRNA amplicons in lane 3.
(b) Fluorescence micrographs of neutrophils labeled with monospecific
anti-RAGE IgG. The cells were viewed by indirect immunofluorescence
using TRITC-conjugated secondary antibody (upper panel). All
images were obtained using confocal microscopy at a vertical resolution
of 2 µm (full width maximum value). Lower panel shows the
presence of RAGE on neutrophil by Western blotting as indicated in
Materials and Methods. The figure shows the absence of RAGE from PNS
and cytosolic fractions and the presence of RAGE in membrane-enriched
fractions of human neutrophils. The numbers at the bottom of the panel
indicate the amount of proteins added in each well (ng).
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Binding AGE albumin to human naïve neutrophils
To test whether the expressed RAGE binds to AGE albumin was next
accomplished by using confocal microscopy of FITC-labeled AGE albumin.
In a series of experiments, FITC-AGE albumin-treated neutrophils
exhibited significantly higher fluorescence intensity than control
protein (FITC albumin). The intensity was significantly reduced in the
presence of soluble RAGE and anti-RAGE antibodies from 88 ± 6.0
to 22 ± 2 and 30 ± 2 absorbance units (AU),
respectively (Fig. 2 a
). Binding FITC-AGE albumin to human neutrophils was further
confirmed by flow cytometry experiments. We found that FITC-AGE albumin
binds to human neutrophils and that the binding was reversed in the
presence of 200-fold, unlabeled AGE albumin (Fig. 2b)
. Because
AGE-modified proteins contain several AGE structures such as CML and
pentosidine, we tested the effect of CML-modified albumin on the
binding of AGE albumin to human neutrophils. We found that iodinated
AGE albumin (125I-AGE albumin) binds selectively to human
neutrophils (Fig. 2c)
, and the binding was inhibited by 200-fold excess
unlabeled AGE albumin or CML-modified albumin. Furthermore, antibodies
to CML totally abolished 125I-AGE albumin-binding to human
neutrophils. This binding was also inhibited by soluble RAGE and
monospecific anti-RAGE IgG (Fig. 2c)
. Binding 125I-AGE
albumin to neutrophils was half-maximal at an AGE-albumin concentration
of 3.7 ± 0.4 nM. The kinetics of this binding fitted a one
binding-site model with a calculated 188 x 103 ± 5.0 x 103 sites per neutrophil (Fig. 2d)
. The
possibility existed that the physical nature of AGE albumin was
different from that of the unmodified protein and that it is this
property that mediates binding to human neutrophils. We tested this by
running samples of AGE albumin and the unmodified protein on PAGE under
reducing and nonreducing conditions. AGE-modified and unmodified
proteins gave one major band of relative molecular weight of 70 kDa
under both conditions. This suggested that the physical nature of AGE
albumin and control albumin was the same and that the binding of AGE
albumin was because of the glycation of the protein rather than the
possible physical structure of the two moieties. Internalization of
FITC-labeled AGE albumin was viewed using confocal microscopy. In a
series of experiments done at room temperature, we found that
extracellular FITC-labeled AGE albumin was internalized in a
time-dependent manner, reaching a maximum at approximately 2 h
(Fig. 2e)
. The internalized FITC-labeled AGE albumin appeared in
membrane-bound vesicles that were stained positive with the lysosomal
marker LysoTracker.

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Figure 2. Binding of AGE albumin to human naïve neutrophils. (a) Upper
panel: Confocal micrographs of binding FITC-AGE albumin to human
neutrophils (i). Incubation with nonglycated albumin resulted in
minimal fluorescence (ii). The binding could be attenuated with the
addition of soluble RAGE (iii) or antibodies against RAGE (iv). (b)
Specific binding of FITC-AGE albumin to human neutrophils as
demonstrated using FACS analysis. Cytofluorograms of human neutrophils
incubated with FITC-AGE albumin, FITC-AGE albumin in the presence of a
200-fold excess of unlabeled AGE albumin, control (unglycated) FITC
albumin, and control FITC albumin in the presence of a 200-fold excess
of unlabeled albumin. The experiments were performed as described in
Materials and Methods. The results are representative of those obtained
from three separate experiments. (c) Inhibition of
125I-AGE-albumin binding to human naïve neutrophils
by soluble RAGE (Sol.RAGE), monospecific anti-RAGE IgG (AntiRAGE),
rabbit anti-CML antibodies (AntiCML), and 200-fold excess of
CML-modified albumin (200 fold CML). The effect of nonimmune rabbit
serum was also shown (NRS). (d) Specific binding of
125I-AGE albumin to human neutrophils. Specific binding was
calculated as the difference between radioactive counts in the absence
(total) and that in the presence of a 200-fold excess of unlabeled AGE
albumin (nonspecific binding). Each point represents the mean ±
SE (n=3), which is representative of five
different independent experiments. (e) Internalization of FITC-AGE
albumin. Fluorescence micrographs at time zero (left) and at 2 h
(2h) following the addition of FITC-AGE albumin (0.36 µM) to the
cells.
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AGE albumin evokes a transient rise in neutrophil calcium
Because a rise in intracellular-free calcium is pivotal to many
neutrophil responses, we investigated the effect of AGEs on calcium
homeostasis. AGE albumin (0.0363.6 µM) caused a dose-dependent
increase in intracellular-free calcium in human neutrophils from an
average resting level of 101 ± 1 nM (n=180 cells) to
248 ± 29 nM in approximately 83 s (Fig. 3a
and b). The response to AGE albumin was
heterogeneous and asynchronous with a dose-dependent increase in the
number of cells, displaying a calcium transient that was at least
twofold higher than the resting levels. At AGE-albumin concentrations
of 0.036, 0.36, and 0.9 µM, the percentage of cells exhibiting a
calcium transient greater than twofold over resting was 37, 62, and
85%, respectively. Control (albumin) protein at the same
concentrations as the AGE albumin did not exhibit a calcium rise. The
ability of AGE albumin to elicit a calcium response in the absence of
extracellular calcium was tested. Cells incubated in Krebs-HEPES buffer
without calcium but with 1 mM ethylenediaminetetraacetate (EDTA) for 5
min prior to stimulation with AGE albumin elicited similar rises in
intracellular-free calcium levels to those obtained in Krebs-HEPES
buffer containing 1.3 mM CaCl2, suggesting that the rise in
intracellular-free calcium level was a result of release from internal
store(s) (unpublished results). This was further confirmed by the
finding that releasing the intracellular calcium stores by pretreatment
of neutrophils with thapsigargin (10 µg/ml) in calcium-free Krebs
containing 1 mM
ethyleneglycol-bis(ß-aminoethylether)-N,N'-tetraacetic
acid (EGTA) abolished the AGE albumin-induced calcium changes. Because
neutrophils contain several different types of calcium stores
[26
], we compared the AGE albumin-releasable store(s) to
the chemotactic-peptide fMLP-releasable store(s). We found that the
extent of the AGE-induced calcium transient was significantly lower
than that elicited by fMLP. At an optimal concentration of fMLP (1
µM) known to have the maximal effect on neutrophil functions, 100%
of the cells responded (n=45) with an average increase in
intracellular-free calcium of 502 ± 37 nM. We also found that
preincubation of human neutrophils with AGE albumin (0.36 µM) for 5
min at room temperature in calcium-free medium decreased the percentage
of responding cells (from 82% to 35%) and the extent of the calcium
rise (from 589 nM to 345 nM; Fig. 3c
) elicited by fMLP (1 µM). In
contrast, pretreatment of cells with fMLP in the absence of
extracellular calcium inhibited further release of calcium by AGE
albumin
AGE albumin induced a transient actin polymerization in human
neutrophils
One of the earliest events following neutrophil activation is a
rapid and transient increase in actin polymerization
[23
]. Therefore, we tested the effect of AGE albumin on
actin polymerization. Using flow cytometry, we found that AGE albumin
(0.36 µM) evoked a transient increase in the binding sites for
rhodamine phallacidin, which is consistent with a transient rise in
actin polymerization. The maximum effect was seen within 510 s and
was followed by a slow decay to prestimulatory levels (Fig. 4 a
). Filamentous-actin (F-actin) levels were still significantly
higher than prestimulatory levels 5 min after activation by AGE
albumin. AGE albumin-induced actin polymerization could be completely
inhibited in the presence of monospecific anti-RAGE IgG (Fig. 4a)
. AGE
albumin-induced actin polymerization was unaffected by inhibition of
the calcium transient with BAPTA-AM and only partly affected by
inhibition of tyrosine phosphorylation by the phosphotyrosine
inhibitor, genistein (unpublished results). Control albumin did not
affect actin polymerization. To confirm the effect of AGE albumin on
actin polymerization, we measured the amount of actin associated with
the Triton X-100-insoluble cytoskeleton by Western blotting and
immunodetection as described previously [27
]. We found
that treatment of neutrophils with AGE albumin (0.36 µM) caused an
increase in the amount of actin associated with the Triton
X-100-insoluble cytoskeleton (Fig. 4b)
. This amount increased with
time, reaching a maximum within 10 s before decaying to
prestimulatory levels. This is consistent with a transient rise in
F-actin.
AGE albumin inhibits transendothelial migration of human
neutrophils
Diapedesis is the process through which neutrophils and other
blood cells leave circulation and enter various tissues in pursuit of
invading organisms or cellular debris. During the process, a cascade of
events is initiated, which culminates in transmigration through the
endothelium and vascular beds. We tested the effect of AGE albumin on
the ability of neutrophils to transmigrate through endothelial cell
monolayers. In a series of experiments, 51Cr-labeled human
neutrophils were allowed to transmigrate through a layer of HAECs. We
found that the ability to transmigrate was impaired in neutrophils that
were pretreated with AGE albumin (15 min) compared with cells
pretreated with control albumin. This inhibition of transendothelial
migration was dose-dependent (Fig. 5 a
) with Ki 0.5 (concentration of AGE albumin that
causes half-maximum inhibition of transmigration) of 0.41 ± 0.07
µM (n=3).

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|
Figure 5. Effect of AGE albumin on transendothelial migration and ROM production
in human naïve neutrophils. (a) Effect of AGE albumin on
transendothelial migration of human neutrophils. Bars indicate the
mean ± SE. This is a representative of four different
experiments with neutrophils isolated from different donors. (b) AGE
albumin-induced potentiation of ROM production in human neutrophils
stimulated with fMLP. Cells were incubated with the indicated
concentration of AGE albumin (in µM) for 1 min prior to addition of
the chemotactic peptide fMLP (1 µM), as indicated by arrows. The
table shows the mean ± SE from three independent
donors. (c) Inhibition of AGE albumin-induced potentiation of ROM
production by the addition of anti-RAGE antibodies (34 µg/ml). Left
trace is ROM-elicited by fMLP alone (arrow, 1 µM). Middle trace is in
the presence of AGE albumin (0.36 µM). Arrow indicates time of fMLP
(1 µM) addition. Arrows in the right trace indicate time of addition
of anti-RAGE IgG (34 µg/ml), AGE albumin (0.36 µM), and fMLP (1
µM), respectively.
|
|
Effect of AGE albumin on reactive oxygen metabolites production
We tested the effect of AGE albumin on neutrophil function by
investigating its ability to modulate the production of ROM. AGE
albumin had no apparent effect on basal neutrophil ROM production.
However, pretreatment of neutrophils with AGE albumin greatly enhanced
the induction of ROM production by fMLP (1 µM). Under such
conditions, AGE albumin enhanced fMLP-induced ROM production in a
concentration-dependent manner with a K1/2 occurring at
0.17 µM and maximum potentiation occurring at 0.36 µM (Fig. 5b)
,
suggesting a pro-oxidant role for AGE albumin. This enhancement was
inhibited in the presence of anti-RAGE IgG (Fig. 5c)
. Control,
nonglycated albumin had no effect on ROM production (unpublished
results).
AGE albumin modifies the phagocytic activity of neutrophils and
protects ingested bacteria from intracellular killing
The effect of AGE albumin on another neutrophil function,
phagocytosis, was investigated using fluorescently labeled heat-killed
S. aureus (HKSA). In a series of experiments, human
neutrophils were pretreated with increasing concentrations of AGE
albumin, and their phagocytic index was measured. These experiments
revealed a dose-dependent increase in the phagocytic index
(K0.5=0.21 µM; Fig. 6 a
). Neutrophil phagocytosis of fluorescent bacteria was unaffected
by pretreatment with a control protein at the same dose. It is
interesting that pretreatment with AGE albumin inhibited HKSA-induced
ROM production. In a series of experiments, AGE albumin was found to
inhibit ROM production by HKSA in a dose-dependent manner with a
Ki 0.5 of 0.46 µM (Fig. 6b)
. Similar results were
obtained using serum-opsonized HKSA. This suggested that although the
number of ingested bacteria was increased in the presence of AGE
albumin, the number of killed bacteria was reduced. To test this
directly, we used live S. aureus in a phagocytic assay.
Neutrophils were exposed to live bacteria for 1 h before counting
the number of ingested live and dead bacteria using the BacLight
live/dead bacteria assay in the presence and absence of AGE albumin at
a concentration that caused maximum inhibition of ROM production (1.1
µM). We found that within 1 h, the average number of bacteria
ingested was 11.3 ± 1.04 and 22.02 ± 0.99
bacteria/neutrophil for control and AGE albumin-treated neutrophils,
respectively. We also found that control neutrophils contained
52.60 ± 3.54% dead bacteria compared with 18.32 ± 1.92%
in AGE albumin-treated neutrophils (P<0.0001; Fig. 6c
).
This was further validated by measurement of the colony-forming ability
of the ingested bacteria. Such measurements revealed that AGE
albumin-treated neutrophils exhibited higher ability to form colonies
on blood agar than albumin-treated cells (Fig. 6d)
. The possibility
that this apparent protection of ingested bacteria by AGE treatment was
a result of interference with phagolysosome fusion was investigated in
cells labeled with the lysosome marker (LysoTracker Red DND) and
FITC-labeled HKSA. Using confocal microscopy, AGE albumin-treated and
control neutrophils exhibited colocalization of the lysosome marker
with the ingested FITC-labeled HKSA, suggesting no apparent difference
in the phagolysosome formation.

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Figure 6. Effect of AGE albumin on phagocytosis. (a) AGE albumin-induced increase
in PI of neutrophils. Experiment was performed as described in
Materials and Methods. Phagocytic index was calculated as the number of
bacteria in neutrophils divided by the number of neutrophils in 50
fields of view. The bar chart represents mean ± SE
(n=3) and is representative of experiments performed on
neutrophils from three different donors. (b) Dose-response effect of
AGE albumin on S. aureus-induced ROM production as measured
by LDCL. The data are representative of three experiments with
neutrophils obtained from three different donors. (c) Effect of AGE
albumin on intracellular killing of ingested, live bacteria. The upper
panel is a confocal micrograph of a representative experiment showing
control neutrophils (albumin) and AGE albumin-treated neutrophils
(AGE-albumin) stained with a BacLight bacteria staining kit so that
dead bacteria appear red, and live bacteria appear green. Histograms
represent the mean ± SE of at least 50 cells. Gray
bars indicate percentage of dead bacteria, and solid bars represent
live bacteria. This is representative of three individual experiments
using neutrophils isolated from three different donors. (d) Colony
formation of ingested, live S. aureus in albumin (1 µM)
and AGE albumin (1 µM).
|
|
 |
DISCUSSION
|
|---|
AGEs have been implicated in the pathogenesis of several diseases
including diabetes [28
29
30
], atherosclerosis
[31
], and Alzheimers [32
,
33
]. Because AGEs accumulate, especially in vascular
walls [3
], recent work has centered on their interaction
with endothelial cells and has led to the isolation and
characterization of a number of binding proteins for AGEs
[3
]. These include AGE-R1, AGE-R2, and AGE-R3,
identified as oligosaccharyltransferase-48 complex (OST-48), protein
kinase substrate 80K-H [34
, 35
], and
carbohydrate-binding protein-35 (CBP-35, galectin-3)
[36
], respectively, and macrophage scavenging receptor
(MSR) [37
], lysozyme and lactoferrin
[38
], and RAGE [3
, 16
]. In
addition, other scavenging receptors have been described
[39
]. Binding of AGEs to lysozyme and lactoferrin
impairs their antibacterial activity [38
], whereas
binding to AGE-R3 has been associated with impaired proinflammatory
responses [40
]. A role for binding AGEs to the
intracellular receptors AGE-R1 and AGE-R2 is not clear, although both
receptors have their own involvement in signal transduction
[39
]. RAGE, the best-characterized cellular receptors
for AGEs [3
, 41
42
43
], is a member of the
immunoglobulin superfamily of cell-surface proteins [3
,
16
] and is expressed in many cell types
[39
]. A number of signaling pathways are activated
following engagement of RAGE. These include mitogen-activated protein
(MAP) kinases, nuclear factor-
B (NF-
B), cdc42/rac, and
p21ras, depending on the cell type [39
,
44
, 45
].
Here, we focus on the direct interaction between AGEs and
polymorphonuclear neutrophilic leukocytes (PMNs), the most abundant
circulating leukocytes and the first line of hosts defense against
invading microorganisms. Because high levels of AGEs have been observed
in many diabetic patients (0.36±0.03 up to 1.13±0.14 µM)
[46
] and because neutrophils from such patients are
dysfunctional [7
8
9
10
11
], the possibility exists that
aberrant neutrophil responses as a result of AGEs might underlie
neutrophil dysfunction in uncontrolled diabetes. We demonstrate the
presence of RAGE at the message level as detected by RT-PCR. The
expression of this message was demonstrated by the positive labeling of
neutrophils with anti-RAGE IgG using indirect immunofluorescence and by
Western blotting experiments, which show the accumulation of RAGE on
enriched neutrophil membrane preparations. Whether the expressed RAGE
binds to AGE albumin was demonstrated by displacement of FITC-labeled
AGE albumin by soluble RAGE and anti-RAGE IgG, as evident by confocal
microscopy and flow cytometry, and by 125I-AGE
albumin-binding studies, which revealed that AGEs bind to human
neutrophils with a Kd of 3.7 ± 0.4 nM
(188x103±5.0x103 sites/cell), which is
different from that shown for macrophages [47
]. Because
it is established that several AGE structures, such as CML, are present
in AGE albumin [30
, 48
], the possibility
existed that binding AGE albumin may be mediated through CML-modified
albumin. This possibility was tested by 125I-AGE
albumin-binding experiments in the presence of antibodies to CML and in
the presence of 200-fold excess CML-modified albumin. Such experiments
revealed the reversal of binding 125I-AGE albumin to human
naïve neutrophils, suggesting that almost all the binding
activities in AGE albumin were because of CML-modified albumin.
In this study, we show that RAGE engagement caused a rapid elevation in
intracellular-free calcium levels in human neutrophils. In keeping with
our previous observations of neutrophil activation [26
],
the calcium response was asynchronous and heterogeneous but
nevertheless dose-dependent over a wide range of AGE concentrations
(0.0363.6 µM). This rise was because of the release of calcium from
an intracellular membrane-enclosed store(s). Evidence for this was
drawn from the finding that removal of extracellular calcium had no
significant effect(s) on the extent or the kinetics of the AGE-induced
calcium transient. At least two distinct types of calcium stores exist
in human neutrophils [49
, 50
]: a
juxtanuclear single-store that appears to be membranous and dispersible
small-storage organelles near the periphery of the cells. Because
stimuli that differentially release one type of calcium stores but not
the other are likely to exhibit differences in transducing signals
following receptor engagement [50
], we studied
interactions between AGE albumin and fMLP-induced calcium signaling.
Pettit and Hallett [51
] have demonstrated that the
chemotactic-peptide fMLP releases the juxtanuclear site, whereas
cross-linking integrins or FcRIIa releases the dispersible sites. The
extent of the calcium rise induced by AGEs was significantly lower
than that elicited by the chemotactic-peptide fMLP (248±29 nM vs.
502±37 nM; P<0.0001). This could be because the
intracellular calcium stores released by the two stimuli are different,
and/or signal processing following AGEs or fMLP insult is different.
The former was tested by prestimulation of calcium release with AGEs in
calcium-free medium (thus releasing calcium stores without
replenishment) followed by stimulation with fMLP to release the
remaining stores. Under such conditions, the percentage of cells
responding to fMLP and the extent of the calcium rise elicited by fMLP
were reduced, suggesting that AGEs and fMLP release calcium from common
stores. This was further confirmed by the finding that releasing the
calcium store with fMLP inhibited any further release of calcium by
AGEs. The ability of AGEs to attenuate the fMLP-induced calcium rise is
noteworthy because it suggests that AGEs may cause aberrant signal
processing with some serious consequences in terms of stimulus-response
coupling. AGE-induced, dose-dependent elevations in intracellular
calcium have also been demonstrated in rat aortic smooth-muscle cells
[52
]. This receptor-mediated increase in intracellular
calcium levels was linked to an increase in muscle-cell proliferation
and could be inhibited by pretreatment with diltiazem, suggesting (at
least in that cell type) the involvement of voltage-gated calcium
channels [53
]. The possibility that the calcium
transient invoked by AGE albumin would interfere with degranulation
and/or receptor up-regulation was tested. We found no evidence of
increased degranulation or receptor up-regulation following
AGE-RAGE-mediated priming. Evidence for this was drawn from
double-labeled experiments using LysoTracker to follow lysosomal
granules and indirect immunofluorescence with anti-RAGE antibodies to
follow receptor up-regulation. Under such conditions, no apparent
difference was observed between control and AGE-treated cells within
the time frame of the experiment (2 h).
Neutrophil receptor-mediated activation by many soluble stimuli
is coupled to a transient actin polymerization [54
,
55
], which is required for chemotactic, phagocytic, and
secretory responses and also for oxidase-activation signaling. We found
that AGE albumin caused a transient increase in actin polymerization as
measured by the increase in rhodamine-phallacidin binding sites and by
an increase in actin associated with the Triton X-100-insoluble
cytoskeleton. The slightly different kinetics observed with the two
methods of measuring actin polymerization is probably because of the
different pools of polymerized actin that each method measures. Neither
calcium transients nor tyrosine phosphorylation appears to be involved
in the AGE-induced actin polymerization. Evidence for this was drawn
from our finding that inhibition of the calcium, transient by previous
treatment with BAPTA-AM and/or inhibition of tyrosine phosphorylation
by genistien, had no apparent effect on AGE-induced actin
polymerization.
One of the major functions of neutrophils is intracellular killing of
phagocytosed microorganisms. This is done by the transient production
of ROM. ROMs are produced as a result of transient activation of the
reduced nicotinamide adenine dinucleotide oxidase system, which reduces
molecular oxygen to superoxide via the monovalent pathway of molecular
oxygen reduction. The oxidase system can be activated by at least two
molecular mechanisms: calcium-dependent and -independent
[56
]. Because AGE albumin caused a transient rise in
intracellular-free calcium, the possibility existed that this might
mediate ROM production. We found that AGEs had no apparent effect on
ROM production when added alone at concentrations up to 3.6 µM,
suggesting that AGE albumin-induced calcium transient was not
sufficient to evoke ROM production. It also suggests that additional
signals essential for ROM production may not be generated following
RAGE engagement. However, fMLP-induced ROM production was enhanced
significantly by AGEs. Whether the AGE-induced calcium transient was
necessary for this enhancement is yet to be delineated. Treatment of
neutrophils with the intracellular calcium chelator BAPTA-AM prior to
AGE-albumin addition inhibited the enhanced fMLP-induced ROM
production. However, this may be attributed to the indiscriminate
chelation of the AGE-induced and the fMLP-induced calcium rise. The
latter is a prerequisite for fMLP-induced ROM production.
A further consequence of AGEs activation of human neutrophils was to
enhance neutrophil phagocytosis. Phagocytosis of unopsonized HKSA was
increased in a dose-dependent manner as measured by the number of
ingested particles in the presence and absence of AGE albumin. This was
not surprising, because we have demonstrated that actin polymerization,
a physiological necessity for phagocytosis, was stimulated in
AGE-treated neutrophils. Surprisingly, however, although AGEs increased
the number of ingested bacteria per cell, ROM production induced by
these bacteria was inhibited in a dose-dependent manner. Furthermore,
experiments with live bacteria revealed that AGEs treatment reduces the
neutrophils ability to kill the ingested bacteria. This coupled with
our finding that FITC-labeled AGE albumin was internalized in vesicles
that colocalized with the lysosomal tracker (LysoTracker Red DND) and
the described finding that AGE albumin directly inhibited lysozymal
activities [38
] suggest that AGE-exposed neutrophils may
themselves contribute to the spread of infection by creating "safe
havens" that promote bacterial evasion of the immune system. This may
serve as the origin of chronic, smoldering, or difficult-to-treat
infections in patients with poorly controlled diabetes and renal
failure. It is noteworthy, however, that studies by Liu et al.
[57
] suggest that AGEs suppress phagocytosis in mouse
peritoneal macrophages and that in a diabetic mouse model, the
decreased phagocytic activity correlated inversely with the AGEs
content of the surrounding tissues. The basis for this apparent
discrepancy is yet to be resolved, but is likely a result of different
cell types studied or species difference or the fact that peritoneal
phagocytic cells differ from peripheral blood phagocytes in many
respects, including maturity.
The effect of AGEs on transendothelial migration is noteworthy. Our
data demonstrate impaired fMLP-induced migration of neutrophils treated
with AGEs in a dose-dependent manner. Many advance glycated proteins
have been shown to exert direct effect on the chemotactic activities of
some cell types [14
, 58
]. Therefore, it is
tempting to speculate that sustained stimulation of neutrophils with
AGEs reduces the cells ability to respond to physiological
chemotactic stimuli. Sengoelge et al. [59
] have shown a
sevenfold increase in transendothelial migration of neutrophils after
exposure of endothelial cells to advanced glycated fibronectin and
inflammatory mediators. It would be interesting to test whether
AGE-treated neutrophils would exhibit a similar increase in
transendothelial migration through endothelial monolayers that have
been exposed to AGE.
It is well-known that in patients with uncontrolled diabetes,
neutrophil function is perturbed [7
8
9
10
11
]. Whether this
perturbation is a consequence of persistent stimulation of the
neutrophils with glycated proteins/lipid is yet to be determined.
Although high levels of glucose directly impair neutrophil function(s)
[60
], AGEs have been shown to activate the vascular
endothelium [61
, 62
], which in turn may
modulate neutrophil functions because the two are in intimate contact.
Therefore, we propose that in diseases where AGE levels are elevated,
inappropriate activation of neutrophils must contribute to the
pathogenesis of that particular disease.
 |
ACKNOWLEDGEMENTS
|
|---|
This work was supported by the Cardiovascular Collaborative
Programme and Research Centre Funds. K. S. C. and R. S. P. contributed equally to this work. We give our gratitude to
Drs. K. Khabar for stimulating discussions and to Dr. K. Al-Hussein and
the Flow Cytometry core facility for their technical help. Our special
thanks go to the staff of the DNA Sequencing core facility.
Received August 25, 2001;
revised October 23, 2001;
accepted November 3, 2001.
 |
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