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Complications

Production of Nε-(Carboxymethyl)Lysine Is Impaired in Mice Deficient in NADPH Oxidase

A Role for Phagocyte-Derived Oxidants in the Formation of Advanced Glycation End Products During Inflammation

  1. Melissa M. Anderson1 and
  2. Jay W. Heinecke2
  1. 1Pharmacia Corporation, St. Louis, Missouri
  2. 2Department of Medicine, University of Washington, Seattle, Washington
  1. Address correspondence and reprint requests to Jay Heinecke, Division of Endocrinology, Metabolism and Nutrition, Box 356426, University of Washington, Seattle, WA 98195. E-mail: heinecke{at}u.washington.edu
Diabetes 2003 Aug; 52(8): 2137-2143. https://doi.org/10.2337/diabetes.52.8.2137
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A Role for Phagocyte-Derived Oxidants in the Formation of Advanced Glycation End Products During Inflammation

Abstract

Advanced glycation end products (AGEs) derived from glucose are implicated in the pathogenesis of diabetic vascular disease. However, many lines of evidence suggest that other pathways also promote AGE formation. One potential mechanism involves oxidants produced by the NADPH oxidase of neutrophils, monocytes, and macrophages. In vitro studies have demonstrated that glycolaldehyde, a product of serine oxidation, reacts with proteins to form Nε-(carboxymethyl)lysine (CML), a chemically well-characterized AGE. We used mice deficient in phagocyte NADPH oxidase (gp91-phox−/−) to explore the role of oxidants in AGE production in isolated neutrophils and intact animals. Activated neutrophils harvested from wild-type mice generated CML on ribonuclease A (RNase A), a model protein, by a pathway that required l-serine. CML formation by gp91-phox−/− neutrophils was impaired, suggesting that oxidants produced by phagocyte NADPH oxidase contribute to the cellular formation of AGEs. To determine whether these observations are physiologically relevant, we used isotope-dilution gas chromatography/mass spectrometry to quantify levels of protein-bound CML in mice suffering from acute peritoneal inflammation. Phagocytes from the gp91-phox−/− mice contained much lower levels of CML than those from the wild-type mice. Therefore, oxidants generated by phagocyte NADPH oxidase may play a role in AGE formation in vivo by a glucose-independent pathway.

  • AGE, advanced glycation end product
  • CGD, chronic granulomatous disease
  • CML, Nε-(carboxymethyl)lysine
  • DTPA, diethylenetriaminepentaacetic acid
  • GC, gas chromatography
  • HOCI, hypochlorus acid
  • HPLC, high-performance liquid chromatography
  • MBTH, 3-methyl-2-benzothiozolinone hydrazone hydrochloride
  • M·−, molecular ion
  • MS, mass spectrometry
  • m/z, mass-to-charge ratio
  • PMA, phorbol myristate acetate
  • RNase A, ribonuclease A

Formation of advanced glycation end products (AGEs) is thought to accelerate the vascular disease seen in diabetes (1–5). One important AGE precursor is glucose, a carbonyl compound present at millimolar concentrations in blood. In vitro, glucose reacts nonenzymatically with amino groups of proteins to form a Schiff base, which undergoes the Amadori rearrangement (1–5). A complex series of oxidative reactions converts the Amadori product to AGEs (1–7). Glycoxidation products affect protein function (1–10) and activate proinflammatory signaling pathways (11), suggesting that they might play a central role in diabetes complications.

Two AGEs, Nε-(carboxymethyl)lysine (CML) and pentosidine, have been well characterized chemically and biologically (6–9,11), but the mechanisms that generate AGEs in vivo are not fully understood. Glucose is implicated because humans with diabetes accumulate CML and pentosidine in skin collagen faster than age- and sex-matched control subjects (7,8). There is also a strong relation between AGE levels and diabetes complications (1–5,7–9). However, AGEs also accumulate at high levels in the plasma proteins of patients with renal failure, regardless of the presence or absence of diabetes (12). This phenomenon has been attributed to increased levels of reactive carbonyls derived from carbohydrate and lipid oxidation (12). Moreover, AGEs have been detected in inflammatory tissue of euglycemic animals and humans (12–14), suggesting that pathways independent of glucose can contribute to AGE formation in vivo.

Reagent glycolaldehyde promotes CML formation in vitro, suggesting that protein-bound glycolaldehyde might be an intermediate in the conversion of glucose to AGEs (15,16). Lipid peroxidation also promotes AGE formation by reactions that might involve reactive carbonyls (17). Moreover, immunohistochemical studies have detected partial colocalization of CML and products of lipid peroxidation in vascular tissue and diabetic kidneys (18). The patterns of immunostaining for CML and pentosidine differ in diabetic tissue, suggesting that different pathways contribute to AGE formation in vivo (18). Collectively, these observations suggest that a complex interplay of oxidative reactions involving reactive carbonyls, lipids, and carbohydrates might increase AGE generation. However, the physiological relevance of these pathways remains to be established.

Another important pathway for generating reactive intermediates involves activated phagocytes (19). These cells use a membrane-associated NADPH oxidase to produce superoxide, which dismutates into hydrogen peroxide (H2O2). Myeloperoxidase, a heme protein secreted by activated phagocytes, uses H2O2 to oxidize chloride ion to hypochlorous acid (HOCl), and the HOCl oxidizes amino acids (20–24). One excellent substrate is l-serine, which becomes glycolaldehyde (21). This aldehyde reacts with model proteins to produce CML in vitro (15,25). Moreover, immunohistochemical studies have detected CML in macrophages of human atherosclerotic lesions (14,26). In contrast with CML levels in skin, however, those in the artery wall are not higher in diabetic subjects than in euglycemic control subjects (14,26). This observation suggests that phagocyte-derived oxidants might be more important than glucose for AGE formation at sites of inflammation, such as atherosclerotic lesions (27).

In the current studies, we used mice deficient in NADPH oxidase (28), the major pathway by which phagocytes generate H2O2, to investigate the role of oxidants in AGE formation in vivo. The animals lack the gp91-phox subunit of the membrane-associated flavocytochrome complex of the oxidase (locus Cybb on the X chromosome), and they recapitulate the clinical features of chronic granulomatous disease (CGD), a genetic disorder that permits recurrent bacterial and fungal infection (19). Our observations strongly suggest that oxidants generated by the NADPH oxidase of phagocytes are important for CML production in vivo, raising the possibility that this pathway plays an important role in AGE formation during inflammation.

RESEARCH DESIGN AND METHODS

Materials.

Crystalline catalase (from bovine liver, thymol-free), ribonuclease A (RNase A; lyophilized from bovine pancreas, code RAF), and glycolaldehyde were purchased from Boehringer Mannheim Biochemicals (Indianapolis, IN), Worthington Biochemical (Freehold, NJ), and Fluka Chemical (Rononkoma, NY), respectively. [d4]CML and CML were provided by Dr. S. Thorpe (University of South Carolina) (8). CGD (Cybb−/−) mice (29) and wild-type (Cybb+/+) littermates in a mixed C57BL/6J and 129-SV genetic background were provided by Dr. R. LeBoeuf (University of Washington). CGD mice (28) and wild-type mice in a C57BL/6J genetic background were provided by Jackson Laboratories (Bar Harbor, MA). All other materials were purchased from Sigma Chemical (St. Louis, MO) unless otherwise indicated.

Isolation of mouse neutrophils.

Mice were injected intraperitoneally with 1 ml 4% sterile thioglycollate broth to recruit neutrophils into the peritoneal cavity (28). Peritoneal inflammatory cells were harvested 16–20 h later by lavage with 10 ml ice-cold medium A (magnesium-, calcium-, phenol-, and bicarbonate-free Hank’s balanced salt solution [Gibco-BRL] supplemented with 100 μmol/l diethylenetriamine pentaacetic acid [DTPA], pH 7.2). DTPA was included in medium A to inhibit metal-catalyzed oxidation reactions. Cells were collected by low-speed centrifugation (400g for 5 min), resuspended in ice-cold medium A, and immediately used for experiments.

Generation of glycolaldehyde by mouse neutrophils.

Neutrophils were isolated from CGD (Cybb−/−) and wild-type (Cybb+/+) mice in the C57BL/6J background (28). Freshly harvested cells (2 × 106/ml) were incubated at 37°C in medium A supplemented with 1 mmol/l l-serine. The cells were stimulated with 200 nmol/l phorbol myristate acetate (PMA) and maintained in suspension by intermittent inversion. The reactions were stopped at the indicated times by pelleting the neutrophils by centrifugation (10 min × 8,000g at 4°C). Supernatants were immediately derivatized and analyzed for glycolaldehyde by high-performance liquid chromatography (HPLC).

Quantifying glycolaldehyde production.

Glycolaldehyde was derivatized with 3-methyl-2-benzothiozolinone hydrazone hydrochloride (MBTH) (30) by incubating 200 μl supernatant for 20 min at room temperature with 568 μl sodium phosphate buffer (5 mmol/l, pH 7.0), 25 μl HCl (6.0 N), and 132 μl MBTH (155 mmol/l). The azine derivative was reacted with 75 μl FeCl3 (370 mmol/l) and incubated for >10 min at room temperature. The resulting blue-colored derivative was stable for at least 5 h. Aldehyde production was quantified by HPLC using a standard curve generated with MBTH derivatives of authentic glycolaldehyde.

HPLC.

Reverse-phase HPLC was performed on a C18 column (5 μm resin, 4.6 × 250 mm, Beckman uPorasil) at a flow rate of 1 ml/min. The column was equilibrated with 70% solvent A (0% methanol and 0.1% trifluoroacetic acid, pH 2.5) and 30% solvent B (100% methanol and 0.1% trifluoroacetic acid, pH 2.5). The column was eluted with a discontinuous gradient of solvent B: 30% solvent B for 3 min, 30–60% solvent B over 2 min, then 60–100% solvent B over 20 min. The eluted MBTH derivatives were detected by absorbance at 598 nm (30).

Reaction conditions for protein modification by mouse neutrophils.

Neutrophils were isolated from CGD (Cybb−/−) and wild-type (Cybb+/+) mice in the C57BL/6J background (28). To modify RNase A, freshly harvested cells (2 × 106 cells/ml) were added to medium B (50 mmol/l sodium phosphate, 100 mmol/l NaCl, 4 mmol/l KCl, and 100 μmol/l DTPA, pH 7.2) supplemented with RNase A (1 mg/ml) and 260 μmol/l l-gluatamate, 210 μmol/l l-alanine, 200 μmol/l l-serine, 175 μmol/l glycine, 165 μmol/l l-valine, 100 μmol/l l-proline, and 100 μmol/l l-lysine (31). Neutrophils were activated with 200 nmol/l PMA and incubated for 60 min at 37°C, with intermittent inversion. After removing the neutrophils by centrifugation (10 min × 8,000g), the mixture was incubated for the indicated amount of time at 37°C. Reactions were terminated by freezing at −20°C. For CML analysis, the samples were dried under vacuum by centrifugal evaporation, 32 pmol [d4]CML and 45 nmol l-[d8]lysine were added, and the protein pellet was hydrolyzed with 6 N HCl for 24 h at 110°C under N2 (32).

Isolation of protein from mouse neutrophils.

Peritoneal inflammatory cells were isolated 24 h after thioglycollate injections from age-matched wild-type (Cybb+/+) and CGD (Cybb−/−) mice in a homogeneous C57BL/6J or mixed C57 BL6/6J and 129-SV genetic background and immediately exposed to 500 mmol/l NaBH4 in 200 mmol/l borate buffer (pH 9) to reduce Schiff bases (32). After an overnight incubation at 4°C, the samples were dialyzed against distilled water for 2 days at 4°C and then delipidated three times using water:methanol:water-washed ether (1:3:7 vol/vol/vol). After the samples were dried by centrifugal evaporation, internal standards were added, and the protein pellet was hydrolyzed with acid as described above.

Analysis of protein-bound CML by negative-ion, chemical ionization gas chromatography/mass spectrometry.

Amino acid hydrolysates were dried in vacuo, resuspended in 1 ml of 1% trifluoroacetic acid, and immediately passed over a solid-phase C18 extraction column (1-ml Supelclean LC-18 SPE tubes; Supelco, Bellefonte, PA) that had been equilibrated with methanol and 1% trifluoroacetic acid. The column was washed with two 1-ml aliquots of 1% trifluoroacetic acid, and the flow-through and two washes were pooled and dried by centrifugal evaporation (32). The n-propyl pentafluoropropionyl derivatives of CML and lysine were prepared and subjected to isotope-dilution gas chromatography/mass spectrometry (GC/MS) as described (25,33). CML, [d4]CML, l-lysine, and l-[d8]lysine were quantified using selected ion monitoring of ions of charge-to-mass ratios (m/z) 560, 564, 460, and 468, respectively, as described (25,32). Selected ion monitoring GC/MS was performed on a Finnigan SSQ equipped with a 12-m DB-1 capillary column (PJ Cobert, St. Louis, MO) (0.2 mm ID, 0.33 μm film thickness). Samples were injected with a 20:1 split with methane as the reagent gas. The injector port and detector were kept at 250°C, with the source at 200°C. To analyze CML and lysine, the initial GC column temperature of 120°C was held for 3 min and then increased from 120 to 300°C at 20°C/min. The temperature was then held at 300°C for 2 min.

RESULTS

Activated neutrophils convert l-serine to glycolaldehyde by a reaction pathway that requires NADPH oxidase and H2O2.

Previous in vitro studies indicate that one mechanism for CML formation in model systems involves the conversion of l-serine into glycolaldehyde by a reaction that is inhibited by catalase, suggesting a requirement for H2O2 (15,21). Mouse neutrophils isolated from wild-type mice and activated with PMA in a balanced salt solution converted l-serine into a compound that reacted with MBTH, a derivatizing agent that forms a stable-colored adduct with aldehydes (30). The derivative comigrated with derivatized authentic glycolaldehyde on reverse-phase HPLC (Fig. 1). Other closely related carbonyl compounds (glyceraldehyde, glyoxal, acrolein, formaldehyde, and acetaldehyde) exhibited HPLC retention times that were distinctly different from that of glycolaldehyde (data not shown). Aldehyde synthesis by the cells required l-serine. In contrast, neutrophils isolated from CGD mice and activated with PMA in the presence of l-serine failed to generate glycolaldehyde (Fig. 1). These observations suggest that conversion of l-serine to glycolaldehyde requires oxidants derived from NADPH oxidase.

When we monitored glycolaldehyde formation by neutrophils isolated from wild-type mice, we observed a short lag phase followed by a linear progress curve that reached a plateau by 90 min (Fig. 2A). Aldehyde production increased with rising concentrations of l-serine (Fig. 2B) and was optimal around neutral pH (Fig. 2C). In contrast, production of free HOCl by myeloperoxidase is optimal under acidic conditions (24), suggesting that other factors affect glycolaldehyde production. Glycolaldehyde formation required activation of the cells with PMA (Fig. 3) and was inhibited by cyanide, implicating a heme protein such as myeloperoxidase in the reaction. Superoxide dismutase (which enhances dismutation of superoxide to H2O2 500-fold at neutral pH) (34) increased the product yield (Fig. 3), perhaps by increasing the availability of H2O2 (35). Alternatively, superoxide dismutase may have prevented superoxide from inactivating myeloperoxidase (36).

In contrast to wild-type neutrophils, neutrophils isolated from CGD mice and activated with PMA failed to convert l-serine into glycolaldehyde under any of the conditions tested (Fig. 3). Moreover, catalase, a peroxide scavenger, inhibited glycolaldehyde production by neutrophils isolated from wild-type mice (Fig. 3). Collectively, these observations indicate that neutrophils use the H2O2 produced by the NADPH oxidase when they generate glycolaldehyde, a reactive α-hydroxyaldehyde that promotes AGE formation in vitro (15).

Phagocytes use oxidants produced by the NADPH oxidase to generate CML on model proteins.

To determine whether activated phagocytes use intermediates derived from NADPH oxidase to produce CML on model proteins, we isolated neutrophils from wild-type mice and then stimulated them with PMA in a physiological salt solution supplemented with RNase A. To mimic a more physiological milieu, plasma concentrations of the seven most common amino acids (260 μmol/l l-gluatamate, 210 μmol/l l-alanine, 200 μmol/l l-serine, 175 μmol/l glycine, 165 μmol/l l-valine, 100 μmol/l l-proline, and 100 μmol/l l-lysine) were included in the medium (31). After a 1-h incubation, the cells were removed by centrifugation and the medium was incubated for 72 h at 37°C. The RNase A was then hydrolyzed with acid, the resulting amino acids were derivatized, and then the derivatives were subjected to analysis by negative-ion electron-capture GC/MS.

The n-propyl pentafluoropropionyl derivative of CML, monitored as ions at m/z 540 (M·− –2 HF) and 560 (M·− –HF), was readily detected in the amino acid mixture by selected ion monitoring (Fig. 4). The relative abundance and retention time of the two most abundant ions derived from the compound generated by the activated neutrophils were identical to those of authentic CML (Fig. 4). During selected ion monitoring, the ions derived from d4-labeled CML eluted slightly earlier than did those of CML, as has been reported for other deuterated internal standards (33). These results indicate that CML is present in acid hydrolysates of a model protein exposed to activated neutrophils.

We next determined the relative importance of oxidants produced by the phagocyte NADPH oxidase to CML production in vitro. CML was quantified in RNase A, which was exposed to PMA-stimulated cells in medium supplemented with plasma concentrations of amino acids using isotope-dilution GC/MS. Neutrophils isolated from wild-type mice markedly increased CML levels (Fig. 5A). Generation of this AGE required stimulation of the neutrophils with PMA, implicating cellular activation and oxidant generation in the reaction (Fig. 5B). Omitting l-serine from the reaction mixture inhibited CML formation by ∼80% (Fig. 5B). Neutrophils isolated from CGD mice produced only low levels of CML (Fig. 5). These results indicate that activated human neutrophils use oxidants generated by their NADPH oxidase to generate CML in a physiological mixture of amino acids. They also indicate that low levels of CML are formed under these conditions by a different pathway independent of NADPH oxidase.

Generation of Nε-(carboxymethyl)lysine during acute inflammation is impaired in CGD mice.

To determine whether oxidants derived from the NADPH oxidase might generate AGEs in vivo, we measured CML levels in inflammatory cells isolated from mice injected intraperitoneally with thioglycollate (Fig. 6). This treatment triggers an inflammatory response that initially recruits neutrophils into the peritoneum (28). Subsequently, the inflammatory cell population evolves into a mixture that contains mostly monocytes and macrophages. This model system therefore mimics many of the cellular events of a classic acute inflammatory response.

Peritoneal inflammatory cells were harvested 24 h after the thioglycollate injections, using wild-type and CGD mice in two genetic backgrounds (age-matched littermates in a mixed 129-SV and C57BL/6J background or age-matched mice in the homogeneous C57BL/6J background). The phagocytes from the C57BL/6J wild-type mice were 78 ± 14% neutrophils, 16 ± 11% macrophages, and 0 ± 1% eosinophils (n = 8). Those from the CGD mice were 76 ± 26% neutrophils, 18 ± 21% macrophages, and 2 ± 2% eosinophils (n = 7). After reduction and delipidation, cellular proteins were hydrolyzed with acid and derivatized. Using isotope-dilution, negative-ion electron-capture GC/MS, we then quantified the n-propyl pentafluoropropionyl derivative of CML. Compared with the samples from the wild-type mice, there was 40% less protein-bound CML in peritoneal inflammatory cells isolated from the CGD mice in both the mixed genetic background (187 ± 53 vs. 109 ± 28 μmol/mol lysine; n = 8 and 5; P < 0.02) and the homogeneous C57BL/J6 background (100 ± 24 vs. 60 ± 8 μmol/mol lysine; n = 7 and 6; P < 0.004). These observations strongly suggest that oxidants derived from phagocyte NADPH oxidase generate CML in vivo.

DISCUSSION

In the current studies, we used animals deficient in NADPH oxidase (28), the major pathway by which phagocytes produce H2O2 (19), to show that oxidants derived from NADPH oxidase contribute to CML formation in vivo. First, we showed that neutrophils isolated from wild-type mice, but not those from mice deficient in NADPH oxidase (CGD mice), converted l-serine into glycolaldehyde. The peroxide scavenger catalase inhibited glycolaldehyde production by wild-type neutrophils. These results indicate that phagocytes generate glycolaldehyde by a reaction dependent on H2O2 produced by the phagocyte NADPH oxidase. Second, CML was produced on RNase A when the protein was exposed to activated wild-type neutrophils in a physiological salt solution, which contained the seven most common plasma amino acids (including l-serine). In contrast, only low levels of CML were formed when we exposed RNase A to neutrophils isolated from the CGD mice. This indicates that oxidants generated by NADPH oxidase are necessary for CML formation under physiologically plausible conditions. Finally, we used isotope-dilution GC/MS analysis, a sensitive and specific analytical method (32,33), to quantify CML in neutrophils isolated from animals with chemically induced peritoneal inflammation. There was markedly less CML in neutrophils isolated from CGD mice than in those isolated from wild-type animals. Collectively, these observations indicate that glucose is not the only pathway for the generation of AGEs and that oxidants derived from phagocyte NADPH oxidase represent one pathway for generating CML in vivo.

Previous studies (8,14,37) have shown that CML increases in skin collagen with aging and that the rate of increase is greater in diabetic subjects than in age- and sex-matched control subjects. In euglycemic humans, CML increases at a rate of ∼4 μmol per mole of lysine per year (8). In mice with chemically induced peritoneal inflammation, we found that wild-type neutrophils had acquired ∼50 μmol more CML per mole of lysine than CGD neutrophils. Because the neutrophils were harvested from the peritoneum 24 h after stimulation, these observations suggest that phagocyte-derived oxidants produce CML during acute inflammation at a rate that is ∼4,000-fold greater than that observed in human skin collagen.

Immunohistochemical studies (13,14,26) have shown that high levels of CML and AGEs are present in the cytoplasm of macrophage foam cells of atherosclerotic lesions in nondiabetic animals and humans. These observations suggest that pathways independent of glucose contribute to CML formation in atherosclerosis, a chronic inflammatory disease. Consistent with this proposal, immunohistochemical studies (14) found little difference between the amount of CML in the macrophages of diabetic and nondiabetic human atherosclerotic lesions. A monoclonal antibody to AGEs that was subsequently shown to react predominantly with CML gave similar results (38). Therefore, an AGE-generating pathway that does not involve glucose might operate in the artery wall. One mechanism may involve myeloperoxidase, which colocalizes in part with lipid-laden macrophages in atherosclerotic lesions (27). Elevated levels of protein and lipid oxidation products characteristic of myeloperoxidase, including an adduct derived from protein-bound glycolaldehyde, have been detected in human atherosclerotic tissue and in LDL isolated from lesions (29,39,40). Another mechanism for accumulating AGEs inside macrophages may involve the uptake of AGE-modified matrix proteins or lipoproteins by cell-surface proteins, such as receptors of advanced glycation end products (RAGEs) or scavenger receptors (11). Importantly, high concentrations of oxidants are secreted into the phagolysosome of macrophages (19), and protein-bound lipid oxidation products are found in lysosomal-like structures in macrophage foam cells (41). These observations indicate that NADPH oxidase might contribute to the formation of CML and other AGEs in macrophage foam cells. In future studies, it will be important to determine whether oxidants produced by phagocytes contribute to AGE formation in humans.

It has been proposed (12) that increased levels of reactive carbonyls—“carbonyl stress”—contribute to AGE formation and the long-term development of complications in diabetes and renal failure. One pathophysiologically relevant mechanism involves glucose, which is a carbonyl in its open chain form. Wolf et al. (42) first proposed that reducing sugars participates in metal ion–dependent oxidation reactions that generate superoxide, H2O2, hydroxyl radicals, and reactive aldehydes. Moreover, oxidation is critical for the formation of CML (1,6). Thus, glycoxidation reactions appear necessary for generating this AGE in vitro. Another potential pathway involves mitochondria, which generate reactive species. Many lines of evidence suggest that glucose increases the production of reactive species inside cultured vascular cells and that inhibitors of mitochondrial electron transport block this increase (43). Lipid oxidation also produces reactive carbonyl intermediates, such as glyoxal, that promote CML formation (16,17). Moreover, epitopes for protein-bound lipid oxidation products and AGEs colocalize in part in atherosclerotic tissue, suggesting that lipid oxidation contributes to AGE formation during atherogenesis (14,17,38). Glucose and lipid oxidation products have also been implicated in the formation of AGEs in renal failure and diabetic nephropathy (1–4,12).

The H2O2 produced by NADPH oxidase might also generate carbonyls and AGEs by other mechanisms. For example, the myeloperoxidase-H2O2 system of phagocytes converts tyrosine to oxidizing intermediates (35). Tyrosyl radical generated by this system peroxidizes lipids by reactions that do not require free metal ions (35). This pathway also generates dityrosine cross-links in proteins, and elevated levels of dityrosine have been detected in human atherosclerotic lesions (44), raising the possibility that carbonyls derived from oxidized lipids promote CML formation in vivo (17). Another potential pathway involves the H2O2-dependent oxidation of protein-bound Amadori and post-Amadori products (45). Reagent H2O2 promotes the formation of CML and pentosidine on glycated collagen in vitro. This pathway may be important physiologically because catalase inhibits the formation of AGEs on collagen in vivo (45). Thus, CML formation at sites of inflammation is likely to involve a complex interplay among oxidants, carbohydrates, lipids, and amino acids.

Our results indicate that oxidants derived from phagocyte NADPH oxidase promote CML formation during acute inflammation in mice. These observations suggest that the generation of reactive intermediates by phagocyte NADPH oxidase contributes to CML formation during atherogenesis. They also support the hypothesis that generation of reactive carbonyls by the oxidation of glucose, lipids, and amino acids is a common mechanism for AGE formation in aging and disease.

FIG. 1.
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FIG. 1.

Reverse-phase HPLC analysis of glycolaldehyde generated by mouse neutrophils. Neutrophils were from the peritoneum of wild-type and CGD mice in a C57BL/6J genetic background. The complete system consisted of neutrophils (2 × 106/ml) suspended in medium A supplemented with 1 mmol/l l-serine (wild-type). When indicated, l-serine was omitted from the medium (wild-type, -serine) or CGD neutrophils were used (CGD). Cells were maintained in suspension by intermittent inversion. After activation with PMA (200 nmol/l), cells were incubated for 60 min at 37°C. The reaction was stopped by removal of the cells by centrifugation. The supernatant was then derivatized with MBTH and subjected to HPLC analysis on a C18 column. Samples were diluted 1:5 in the derivatization procedure, and the HPLC injection volume was 100 μl. The arrow indicates the retention time of the MBTH derivative of authentic glycolaldehyde.

FIG. 2.
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FIG. 2.

Reaction requirements for glycolaldehyde generation by mouse neutrophils. Neutrophils were isolated from the peritoneum of wild-type mice in a C57BL/6J genetic background. The complete system consisted of PMA-stimulated neutrophils (2 × 106/ml) incubated for 60 min at 37°C in medium A supplemented with 1 mmol/l l-serine as described in the legend to Fig. 1. The cells were incubated for varying time periods (A), with varying concentrations of l-serine (B), or at different pH levels (C). Glycolaldehyde production was quantified by HPLC after the medium was derivatized with MBTH. Data are the mean of four determinations from two independent experiments.

FIG. 3.
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FIG. 3.

Generation of glycolaldehyde by neutrophils isolated from wild-type and CGD mice. Neutrophils were isolated from wild-type and CGD mice in a C57BL/6J genetic background. The complete system (Complete) consisted of PMA-stimulated neutrophils (2 × 106/ml) incubated for 60 min at 37°C in medium A supplemented with 1 mmol/l l-serine as described in the legend to Fig. 1. When indicated, superoxide dismutase (170 nmol/l), catalase (400 nmol/l), or sodium cyanide (10 mmol/l) was included, or l-serine, PMA, or cells were omitted from the complete reaction mixture. The reaction was stopped by removing the cells by centrifugation. The concentration of the MBTH derivative of glycolaldehyde in the supernatant was determined by HPLC analysis. Data are the mean and SE of six determinations from two independent experiments.

FIG. 4.
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FIG. 4.

Detection of the n-propyl pentafluoropropionyl derivative of CML generated by mouse neutrophils by negative-ion electron-capture GC/MS analysis with selected ion monitoring. Wild-type mouse neutrophils (2 × 106/ml) in a C57BL/6J genetic background were incubated at 37°C in medium B supplemented with 1 mg/ml RNase A, 260 μmol/l l-gluatamate, 210 μmol/l l-alanine, 200 μmol/l l-serine, 175 μmol/l -glycine, 165 μmol/l l-valine, 100 μmol/l l-proline, and 100 μmol/l l-lysine. Cells were stimulated with 200 nmol/l PMA, maintained in suspension by intermittent inversion, and removed by centrifugation after 60 min. The medium was then incubated for 72 h. Protein-bound CML in the incubation medium was detected by negative-ion electron-capture GC/MS analysis with selected ion monitoring following reduction, acid hydrolysis, and derivatization of the reisolated amino acids. The ions at m/z 560 (M·− –HF) and 540 (M·− –2 HF) represent the most abundant ions in the full-scan mass spectrum of CML. Note that the ions derived from d4-labeled CML at m/z 564 and 544 elute slightly earlier than the corresponding ions of CML (arrow), as has been shown for many other deuterated compounds (33).

FIG. 5.
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FIG. 5.

Progress curve and reaction requirements for protein-bound CML formation by wild-type and CGD mouse neutrophils. Neutrophils were isolated from wild-type and CGD mice in a C57BL/6J genetic background. The reaction mixture consisted of PMA-stimulated mouse neutrophils (2 × 106/ml) incubated at 37°C in medium B supplemented with 1 mg/ml RNase A and amino acids as described in the legend to Fig. 4. Where indicated, PMA or l-serine was omitted from the reaction mixture. The medium of the cells was analyzed for protein-bound CML at the indicated time (A) or after 72 h incubation (B) by isotope dilution negative-ion electron-capture GC/MS analysis with selected ion monitoring. Data are the mean and SE of four determinations from two independent experiments and are corrected for the endogenous CML content (46.5 or 24.1 μmol/mol) of RNase A. A: Two-way ANOVA revealed that time was a significant variable (P = 0.0006, F = 11.2, and df = 3), that cell type (wild-type versus CGD) was a significant variable (P = 0.003, F = 5.2, and df = 1), and that the interaction between the two factors was significant (P = 0.02, F = 3.9, and df = 3). B: One-way ANOVA with a Dunnett’s post hoc test for planned comparisons revealed that CML production was significantly different for wild-type cells in complete medium (Complete) versus CGD cells in complete medium (P < 0.01) and for wild-type cells in complete medium versus wild-type cells in medium lacking serine (-serine) or medium lacking PMA (-PMA) (P < 0.01).

FIG. 6.
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FIG. 6.

Isotope-dilution GC/MS quantification of protein-bound CML in peritoneal inflammatory cells isolated from wild-type and CGD mice. Inflammatory cells were isolated from age-matched CGD and wild-type mice 24 h after intraperitoneal injection of thioglycollate. After reduction and delipidation, cellular proteins were hydrolyzed and derivatized. The n-propyl pentafluoropropionyl derivative of CML in the amino acid hydrolysate was quantified by isotope-dilution, negative-ion electron-capture GC/MS. A: Cells isolated from littermates of mice with a 129-SV and C57BL/J6 genetic background. B: Cells isolated from mice with a homogeneous (>10 backcrosses) C57BL/J6 genetic background.

Acknowledgments

This work was supported by National Institutes of Health Grants DK02456 and AG021191.

We thank Dr. Susan Thorpe (University of South Carolina) for providing natural-abundance and isotope-labeled CML, Dr. Rene LeBoeuf (University of Washington) for providing CGD mice, and Jennifer Moreno for excellent technical assistance.

Footnotes

    • Accepted May 12, 2003.
    • Received April 15, 2003.
  • DIABETES

REFERENCES

  1. ↵
    Baynes JW: Role of oxidative stress in development of complications in diabetes. Diabetes40 :405 –412,1991
    OpenUrlAbstract/FREE Full Text
  2. Brownlee M, Cerami A, Vlassara H: Advanced glycosylation end products in tissue and the biochemical basis of diabetic complications. N Engl J Med318 :1315 –1321,1988
    OpenUrlCrossRefPubMedWeb of Science
  3. Monnier VM, Sell DR, Nagaraj RH, Miyata S: Mechanisms of protection against damage mediated by the Maillard reaction in aging. Gerontology37 :152 –165,1991
    OpenUrlCrossRefPubMedWeb of Science
  4. ↵
    Brownlee M: Biochemistry and molecular cell biology of diabetic complications. Nature414 :813 –820,2001
    OpenUrlCrossRefPubMedWeb of Science
  5. ↵
    Ledl F, Schleicher E: New aspects of the Maillard reaction in foods and in the human body. Angew Chem Int Ed Engl29 :565 –594,1990
    OpenUrlCrossRef
  6. ↵
    Ahmed M, Thorpe SR, Baynes JW: Identification of Nε-(carboxymethyl) lysine as a degradation product of fructoselysine in glycated proteins. J Biol Chem61 :4889 –4894,1986
    OpenUrl
  7. ↵
    Sell DR, Monnier VM: Structure elucidation of a senescence cross-link from human extracellular matrix: implications of pentoses in the aging process. J Biol Chem264 :21597 –21602,1989
    OpenUrlAbstract/FREE Full Text
  8. ↵
    Dyer DG, Dunn JA, Thorpe SR, Bailie KE, Lyons TJ, McCance DR, Baynes JW: Accumulation of Maillard reaction products in skin collagen in diabetes and aging. J Clin Invest91 :2463 –2469,1993
  9. ↵
    Monnier VM, Glomb M, Elgawish A, Sell DR: The mechanism of collagen cross-linking in diabetes: a puzzle nearing resolution. Diabetes45 (Suppl. 3) :S67 –S72,1996
  10. ↵
    Maillard LC: Action des acides amines sur les sucres: formation des melaniodines par voie methodique. CR Acad Sci154 :66 –68,1912
    OpenUrl
  11. ↵
    Kislinger T, Fu C, Huber B, Qu W, Taguchi A, Du Yan S, Hofmann M, Yan SF, Pischetsrieder M, Stern D, Schmidt AM: N(epsilon)-(carboxymethyl)lysine adducts of proteins are ligands for receptor for advances glycation end products that activate cell signaling pathways and modulate gene expression. J Biol Chem274 :31740 –31749,1999
    OpenUrlAbstract/FREE Full Text
  12. ↵
    Miyata T, Van Ypersele De Strihou C, Kurokaka K, Baynes JW: Alterations in nonenzymatic biochemistry in uremia: origin and significance of “carbonyl stress” in long-term uremic complications. Kidney Internatl55 :389 –399,1999
    OpenUrlCrossRefPubMedWeb of Science
  13. ↵
    Palinski W, Koschinsky T, Butler SW, Miller E, Vlassara H, Cerami A, Witztum JL: Immunological evidence for the presence of advanced glycation end products in atherosclerotic lesions of euglycemic rabbits. Arterio Thromb Vasc Wall Biol15 :571 –582,1995
    OpenUrl
  14. ↵
    Schleicher ED, Wagner E, Nerlich AG: Increased accumulation of the glycoxidation product N(epsilon)-(carboxymethyl)lysine in human tissues in diabetes and aging. J Clin Invest99 :457 –468,1997
    OpenUrlCrossRefPubMedWeb of Science
  15. ↵
    Glomb MA, Monnier VM: Mechanism of protein modification by glyoxal and glycolaldehyde, reactive intermediates of the Maillard reaction. J Biol Chem270 :10017 –10026,1995
    OpenUrlAbstract/FREE Full Text
  16. ↵
    Farboud B, Aotaki-Keen A, Miyata T, Hjelmeland LM, Handa JT: Development of a polyclonal antibody with broad epitope specificity for advanced glycation end products and localization of these epitopes in Bruch’s membrane of the aging eye. Mol Vis14 :5 –11,1999
    OpenUrl
  17. ↵
    Fu M, Requena JR, Jenkins AJ, Lyons TJ, Baynes JW, Thorpe SR: The advanced glycation product Nε-(carboxymethyl)lysine, is a product of both lipid peroxidation and glycoxidation reactions. J Biol Chem271 :9982 –9986,1996
    OpenUrlAbstract/FREE Full Text
  18. ↵
    Horie K, Miyata T, Maeda K, Miyata S, Sugiyama S, Sakai H, van Ypersole de Strihou C, Monnier VM, Witztum JL, Kurokawa K: Immunohistochemical colocalization of glycoxidation products and lipid peroxidation products in diabetic renal glomerular lesions: implication for glycoxidative stress in the pathogenesis of diabetic nephropathy. J Clin Invest100 :2995 –3004,1997
    OpenUrlCrossRefPubMedWeb of Science
  19. ↵
    Babior BM, Lambeth JD, Nauseef W: The neutrophil NADPH oxidase. Arch Biochem Biophys397 :342 –344,2002
    OpenUrlCrossRefPubMedWeb of Science
  20. ↵
    Hazen SL, Hsu FF, Heinecke JW: p-Hydroxyphenylacetaldehyde is the major product of L-tyrosine oxidation by activated human phagocytes: a chloride-dependent mechanism for the conversion of free amino acids into reactive aldehydes by myeloperoxidase. J Biol Chem271 :1861 –1867,1996
    OpenUrlAbstract/FREE Full Text
  21. ↵
    Anderson MM, Hazen SL, Hsu FF, Heinecke JW: Human neutrophils employ the myeloperoxidase-hydrogen peroxide-chloride system to convert hydroxy-amino acids into glycolaldehyde, 2-hydroxypropanal, and acrolein. J Clin Invest99 :424 –432,1997
    OpenUrlCrossRefPubMedWeb of Science
  22. Zgliczynski JM, Stelmaszynska T, Domanski J, Ostrowski W: Chloramines as intermediates of oxidation reactions of amino acids by myeloperoxidase. Biochim Biophys Acta235 :419 –424,1971
    OpenUrlPubMed
  23. Strauss RR, Paul BB, Jacobs AA, Sbarra AJ: Role of the phagocyte in host-parasite interactions–Myeloperoxidase-H2O2-Cl−-mediated aldehyde formation and its relationship to antimicrobial activity. Infec Immun3 :595 –602,1971
    OpenUrlAbstract/FREE Full Text
  24. ↵
    Harrison JE, Schultz J: Studies on the chlorinating activity of myeloperoxidase. J Biol Chem251 :1371 –1374,1976
    OpenUrlAbstract/FREE Full Text
  25. ↵
    Anderson MM, Requena JR, Crowley JR, Thorpe SR, Heinecke JW: The myeloperoxidase system of human phagocytes generates Nε-(carboxymethyl)lysine on proteins: a mechanism for producing advanced glycation end products at sites of inflammation. J Clin Invest104 :103 –113,1999
    OpenUrlCrossRefPubMedWeb of Science
  26. ↵
    Sakata N, Imanaga Y, Meng J, Tachikawa Y, Takebayashi S, Nagai R, Horiuchi S, Itabe H, Takano T: Immunohistochemical localization of different epitopes of advanced glycation end products in human atherosclerotic lesions. Atherosclerosis141 :61 –75,1998
    OpenUrlCrossRefPubMedWeb of Science
  27. ↵
    Daugherty A, Dunn JL, Rateri DL, Heinecke JW: Myeloperoxidase, a catalyst for lipoprotein oxidation, is expressed in human atherosclerotic lesions. J Clin Invest94 :437 –444,1994
  28. ↵
    Pollock JD, Williams DA, Gifford MA, Li LL, Du X, Fisherman J, Orkin SH, Doerschuk CM, Dinauer MC: Mouse model of X-linked chronic granulomatous disease, an inherited defect in phagocyte superoxide production. Nat Genet9 :202 –209,1995
    OpenUrlCrossRefPubMedWeb of Science
  29. ↵
    Hazen SL, Heinecke JW: 3-Chlorotyrosine, a specific marker of myeloperoxidase-catalyzed oxidation, is markedly elevated in low density lipoprotein isolated from human atherosclerotic intima. J Clin Invest99 :2075 –2081,1997
    OpenUrlCrossRefPubMedWeb of Science
  30. ↵
    Paz MA, Blumenfeld OO, Rojkind M, Henson E, Furfine C, Gallop PM: Determination of carbonyl compounds with N-methyl benzothiazolone hydrazone. Arch Biochem Biophys109 :548 –559,1965
  31. ↵
    Linder MC: Nutritional Biochemistry and Metabolism. New York, Elsevier,1992 , p.98
  32. ↵
    Knecht KJ, Dunn JA, McFarland KF, McCance DR, Lyons TJ, Thorpe SR, Baynes JW: Effect of diabetes and aging on carboxymethyllysine levels in human urine. Diabetes40 :190 –196,1991
    OpenUrlAbstract/FREE Full Text
  33. ↵
    Heinecke JW, Hsu FF, Crowley JR, Hazen SL, Leeuwenburgh C, Mueller DM, Rasmussen JE, Turk J: Detecting oxidative modification of biomolecules with isotope dilution mass spectrometry: sensitive and quantitative assays for oxidized amino acids in proteins and tissues. Meth Enzym300 :124 –144,1998
    OpenUrl
  34. ↵
    Babior BM, Kipnes RS, Curnutte JT: Biological defense mechanisms: the production by leukocytes of superoxide, a potential bactericidal agent. J Clin Invest52 :741 –744,1973
  35. ↵
    Savenkova MI, Mueller DM, Heinecke JW: Tyrosyl radical generated by myeloperoxidase is a physiological catalyst for initiation of lipid peroxidation in low density lipoprotein. J Biol Chem269 :20394 –20400,1994
    OpenUrlAbstract/FREE Full Text
  36. ↵
    Kettle AJ, Sangster DF, Gebicki JM, Winterbourn C: A pulse radiolysis investigation of the reactions of myeloperoxidase with superoxide and hydrogen peroxide. Biochim Biophys Acta956 :58 –62,1988
    OpenUrlCrossRefPubMed
  37. ↵
    McCance DR, Dyer DG, Dunn JA, Bailie KE, Thorpe SR, Baynes JW, Lyons TJ: Maillard reaction products and their relation to complications in insulin-dependent diabetes mellitus. J Clin Invest91 :2470 –2478,1993
  38. ↵
    Kume S, Takeya M, Mori T, Araki N, Suzuki H, Horiuchi S, Kadama T, Miyauchi Y, Takahashi K: Immunohistochemical and ultrastructural detection of advances glycation end products in atherosclerotic lesions of human aorta with a novel specific monoclonal antibody. Am J Pathol147 :654 –667,1995
    OpenUrlPubMedWeb of Science
  39. ↵
    Heller JI, Crowley JR, Hazen SL, Salvay DM, Wagner P, Pennathur S, Heinecke JW: p-Hydroxyphenylacetaldehyde, an aldehyde generated by myeloperoxidase, modifies phospholipid amino groups of low density lipoprotein in human atherosclerotic intima. J Biol Chem275 :9957 –9962,2000
    OpenUrlAbstract/FREE Full Text
  40. ↵
    Nagai R, Hayashi CM, Xia L, Takeya M, Horiuchi S: Identification in human atherosclerotic lesions of GA-pyridine, a novel structure derived from glycolaldehyde-modified proteins. J Biol Chem277 :48905 –48912,2002
    OpenUrlAbstract/FREE Full Text
  41. ↵
    Rosenfeld ME, Palinski W, Yla-Herttuala S, Butler S, Witztum JL: Distribution of oxidation specific lipid-protein adducts and apolipoprotein B in atherosclerotic lesions of varying severity from WHHL rabbits. Arteriosclerosis10 :336 –349,1990
    OpenUrlAbstract/FREE Full Text
  42. ↵
    Wolff SP, Jiang ZY, Hunt JW: Protein glycation and oxidative stress in diabetes mellitus and aging. Free Rad Biol & Med10 :339 –352,1991
    OpenUrlCrossRefPubMedWeb of Science
  43. ↵
    Brownlee M: Biochemistry and molecular cell biology of diabetic complications. Nature414 :813 –820,2001
  44. ↵
    Leeuwenburgh C, Rasmussen JE, Hsu FF, Mueller DM, Pennathur S, Heinecke JW: Mass spectrometric quantification of markers for protein oxidation by tyrosyl radical, copper, and hydroxyl radical in low density lipoprotein isolated from human atherosclerotic plaques. J Biol Chem272 :3520 –3526,1997
    OpenUrlAbstract/FREE Full Text
  45. ↵
    Elgawish A, Glomb M, Friedlander M, Monnier VM: Involvement of hydrogen peroxide in collagen cross-linking by high glucose in vitro and in vivo. J Biol Chem271 :12964 –12971,1996
    OpenUrlAbstract/FREE Full Text
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Production of Nε-(Carboxymethyl)Lysine Is Impaired in Mice Deficient in NADPH Oxidase
Melissa M. Anderson, Jay W. Heinecke
Diabetes Aug 2003, 52 (8) 2137-2143; DOI: 10.2337/diabetes.52.8.2137

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Production of Nε-(Carboxymethyl)Lysine Is Impaired in Mice Deficient in NADPH Oxidase
Melissa M. Anderson, Jay W. Heinecke
Diabetes Aug 2003, 52 (8) 2137-2143; DOI: 10.2337/diabetes.52.8.2137
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