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(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{dagger}, David M. Stern{dagger} and Futwan A. Al-Mohanna*

* Biological & Medical Research, King Faisal Specialist Hospital and Research Centre, Riyadh, Saudi Arabia; and
{dagger} 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
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
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


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
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 body’s immune defense, resulting in delayed wound healing [6 ] and an increased susceptibility to infection [7 ].

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.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
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 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){alpha} levels in the conditioned medium were measured using enzyme-linked immunosorbent assay (ELISA; R&D Systems, Minneapolis, MN.). IL-1{alpha} 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.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
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 98–99% pure [as detected by fluorescein-activated cell sorter (FACS) analysis] with 1–2% 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).

 
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.

 
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 (0.036–3.6 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



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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; {blacklozenge}, 0.36 µM, {blacktriangleup}, 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 ({blacksquare}), after preincubation with AGE albumin (0.36 µM, {blacktriangleup}), or with AGE albumin alone ({blacklozenge}) 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.

 
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 5–10 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.



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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 ({blacklozenge}). Actin polymerization was blocked completely by pretreatment with anti-RAGE antibodies ({blacksquare}). (b) Transient increase in F-actin associated with the Triton X-100-insoluble cytoskeleton sampled at the time intervals indicated.

 
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
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
AGEs have been implicated in the pathogenesis of several diseases including diabetes [28 29 30 ], atherosclerosis [31 ], and Alzheimer’s [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-{kappa}B (NF-{kappa}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.


   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.


   REFERENCES
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 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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