* 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
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Key Words: calcium • ROM • transmigration • phagocytosis • signal
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Neutrophils, the predominant circulating leukocytes, are the body’s 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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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 2–10 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 manufacturer’s
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
manufacturer’s protocol. Sense and antisense primers for human RAGE
were 5'-AGCGGCTGGAATGGAAACTGAA-3' (M91211; nucleotides 140–162) and
5'-CTACAGGAGAAGGTGGGACGGG-3' (M1211; reverse complement of nucleotides
605–627), 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 manufacturer’s 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 12–18 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 manufacturer’s 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.
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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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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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Figure 3. Calcium mobilization in AGE-activated human neutrophils. (a)
Dose-dependent increase in calcium response to AGE albumin. Neutrophils
were loaded with the fluorescent calcium indicator Fura 2-AM as
described in Materials and Methods. The graph represents calcium
response to various doses of AGE albumin (1.8 µM •, 0.9 µM;  ,
0.36 µM,  , 0.036 µM+) over time (s). (b) Intracellular calcium
maps before (0s) and at the indicated time intervals following
AGE-albumin stimulation. Calcium changes are color-coded (color bar) so
that high calcium concentrations appear hot. This is a representative
experiment obtained from seven different donors. (c) Activation of
Ca2+ transient by fMLP alone ( ), after
preincubation with AGE albumin (0.36 µM, ), or with AGE albumin
alone () in the absence of extracellular calcium. Adherent
neutrophils were preincubated for 5 min in Krebs-HEPES buffer without
CaCl2 but containing EGTA as described in Materials and
Methods.
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Figure 4. Effect of AGE albumin on actin polymerization. (a) Transient increase
in F-actin as measured by flow cytometry of AGE albumin-stimulated,
rhodamine phallacidin-stained neutrophils (). Actin polymerization
was blocked completely by pretreatment with anti-RAGE antibodies ().
(b) Transient increase in F-actin associated with the Triton
X-100-insoluble cytoskeleton sampled at the time intervals indicated.
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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.
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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).
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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 host’s 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 (0.036–3.6 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.
Received August 25, 2001; revised October 23, 2001; accepted November 3, 2001.
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-(carboxymethyl)lysine and N-(carboxymethyl)hydroxylysine in human skin collagen Biochemistry 30,1205-1210[Medline]B require the cytoplasmic domain of the receptor but different downstream signaling pathways J. Biol. Chem. 274,19919-19924- (carboxymethyl)lysine adducts of proteins are ligands for receptor for advanced glycation end products that activate cell signaling pathways and modulate gene expression J. Biol. Chem. 44,31740-31749This article has been cited by other articles:
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