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The Journal of Lipid Research, Vol. 39, 2201-2208, November 1998
Copyright © 1998 by Lipid Research, Inc.

Original Article

LDL oxidation by activated monocytes: characterization of the oxidized LDL and requirement for transition metal ions

Correspondence to: Carl P. Sparrow.

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

Monocytes can be activated by incubation with opsonized zymosan (Zop), and under these conditions can oxidize low density lipoprotein (LDL). We have characterized the biochemical changes in the lipoprotein after this oxidation. We found that monocyte-oxidized LDL has increased mobility on agarose gels, increased absorbance at 234 nm, increased content of lysophosphatidylcholine, and fluorescence at 430 nm when excited at 350 nm. All these features were somewhat less pronounced in monocyte-oxidized LDL than in LDL oxidized by 5 µM CuSO₄. Under appropriate conditions, Zop-stimulated monocytes oxidized LDL to a form recognized by macrophage scavenger receptors. Monocytes stimulated by Zop produced superoxide and also oxidized LDL, whereas monocytes stimulated by phorbol ester produced slightly more superoxide but did not oxidize LDL. We found that the chelators EDTA and diethylenetriaminepentaacetic acid inhibited LDL oxidation by Zop-stimulated monocytes, implying a requirement for transition metal ions. We found that Zop contained approximately 5 nmol iron per mg, probably as Fe³⁺. Zop stripped of its iron supported superoxide production by monocytes, but did not support LDL oxidation. Furthermore, Fe²⁺ appeared in the medium when monocytes were incubated with Zop, but not with iron-stripped Zop.

Taken together, these results imply that monocytes stimulated by Zop are able to oxidize LDL only because of contaminating iron in the commercial zymosan preparations.—Xing, X., J. Baffic, and C. P. Sparrow. LDL oxidation by activated monocytes: characterization of the oxidized LDL and requirement for transition metal ions. J. Lipid. Res. 1998. 39: 2201–2208.

Supplementary key words: scavenger receptor, atherosclerosis, iron, cholesterol, zymosan, superoxide

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

Low density lipoprotein (LDL) oxidation is believed to be an important feature of the biochemical chain of events leading from elevated cholesterol levels to atherosclerosis (1) (2). Oxidized LDL can be detected in atherosclerotic arterial tissue (3), and certain antioxidants slow the progression of atherosclerosis in animal models (4) (5) (6) (7). The importance of oxidized LDL in vascular disease has led to significant research efforts on the causes and effects of LDL oxidation.

LDL oxidation is a complex process. The initial event is probably formation of hydroperoxides in the polyunsaturated fatty acyl chains of LDL lipid(s). Lipid hydroperoxides are conjugated dienes, and their absorbance at 234 nm allows for a simple assay for the early stages of LDL oxidation (8). The hydroperoxides in LDL can decompose to form other oxidizing species, causing a free radical chain reaction within the particle or perhaps between LDL particles. This uncontrolled lipid peroxidation yields many different molecular species (9), some of which can be detected by the thiobarbituric acid assay (10). LDL phospholipids with an oxidized fatty acyl chain at C-2 are substrates for a phospholipase A ₂ that converts them to lysolipids (11). The LDL apolipoprotein (apo)B is altered in two ways by oxidation: there is direct oxidative scission of peptide bonds (12), and some of the products of lipid peroxidation react with the free amino groups of apoB, forming covalent adducts (6) (13). These adducts increase the net negative charge of the particle, which can be easily detected by agarose gel electrophoresis (14), and these adducts also have a characteristic fluorescence spectrum (13) (15). Most importantly, it is the presence of these adducts that causes recognition of oxidized LDL by the scavenger receptor of macrophages (15), and macrophage uptake of oxidized LDL may lead to foam cell formation in vivo (1).

Most characterizations of oxidized LDL are from in vitro studies. LDL can be oxidatively modified in vitro by various chemical agents and by certain cultured cell types (reviewed in refs. 1 and 16). To the extent that LDL oxidation occurs in vivo, it is presumably catalyzed by the cells of the artery. Because of the pathophysiological importance of oxidized LDL, it may be useful to understand the mechanism(s) by which cells oxidize LDL.

LDL oxidation by most cultured cells requires a medium containing transition metal ions (17) (18) (19) (20) (21) (22) (23) (24). These observations present a puzzle: if LDL oxidation requires free transition metal ions, how can LDL by oxidized in vivo given the normal presence of physiological chelators? Perhaps the in vitro systems that require transition metal ions do not accurately mimic the in vivo situation. It would be interesting to find a cellular mechanism for LDL oxidation that does not require transition metal ions. Monocytes activated with opsonized zymosan (Zop) can oxidize LDL in RPMI medium, which is formulated without transition metal ions (25) (26). This intriguing observation implies that LDL may be oxidized in the absence of metal ions by activated monocyte/macrophages in the sub-endothelial space.

Given the potential importance of transition metal-independent LDL oxidation, it would be of great interest to understand the mechanism of LDL oxidation by activated monocytes in RPMI medium. It has been demonstrated that LDL oxidation in this system requires cytosolic phospholipase A ₂ (27) and protein kinase C (28) but does not require lipoprotein-receptor interactions (29). In the present work, we have characterized LDL oxidation by activated monocytes in RPMI medium: the monocyte-oxidized LDL has been analyzed biochemically, and mechanistic studies have addressed the roles of superoxide and transition metal ions.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

Cells and cell culture
Human peripheral blood monocytes were isolated from heparinized blood using Histopaque-1077 as recommended by the supplier (Sigma, St. Louis, MO). The mononuclear cells were plated into cell culture-treated plastic 35-mm wells (5 million cells per well in 6-well plates to yield 1 million monocytes per well). The monocytes were allowed to adhere for 2 h, and then the non-adherent lymphocytes were removed by washing. This procedure yields a relatively pure culture of monocytes (30). The cells were either used immediately or cultured overnight (no significant differences were seen between immediate or overnight use). HL-60 cells, a human monocyte-like cell line, were obtained from ATCC (Rockville, MD) and grown as recommended. The rabbit aortic endothelial cell line RECB4 (31) was grown as previously described (32). Resident mouse peritoneal macrophages were obtained from female Swiss-Webster mice by peritoneal lavage with phosphate-buffered saline. Macrophages were plated in 6-well cell culture plates at 1 million cells/well (for LDL oxidation) or in 24-well plates at 0.6 million cells/well (for uptake assays) in DMEM containing 10% fetal bovine serum.

Lipoprotein
Plasma was obtained from fasted normal volunteers, and LDL were isolated by standard procedures (33). Concentrations of LDL are expressed on the basis of protein, measured using the micro BCA (bicinchoninic acid) protein reagent (Pierce Chemical Co., Rockford, IL), with bovine serum albumin as the standard. LDL was iodinated with Na¹²⁵I using the "trapped label" tyramine cellobiose (34). No significant differences were found in preliminary experiments comparing the oxidative modification of LDL labeled with ¹²⁵I-tyramine cellobiose and LDL labeled conventionally (data not shown).

Preparation of opsonized zymosan (Zop) and stripping of iron from zymosan
Zymosan was purchased from Sigma (St. Louis) and United States Biochemical (Cleveland); similar results were obtained with zymosan from the two different sources. Zymosan was opsonized using human serum as described (30). During the course of these studies, we considered the possibility that commercial zymosan preparations contained Fe³⁺. We therefore developed a method to simultaneously strip zymosan of iron and quantitate the iron removed. Our method used ascorbate to reduce Fe ³⁺ to Fe²⁺, and o-phenanthroline to chromogenically bind Fe ²⁺ (35). Specifically, iron was removed from zymosan by suspending 0.5 g zymosan in 40 ml of stripping solution, which contained 50 mM HEPES (pH 7.4), 1 mM diethylenetriaminepentaacetic acid (DETAPAC), 100 µM o-phenanthroline, and 10 mM ascorbate (all solutions used were sterile filtered). The suspension, in a sterile 50-ml conical tube, was rotated slowly at room temperature for 2–3 days with 4 or 5 changes of stripping solution. The last two treatments also contained 20% ethanol. Zymosan is particulate, so the material can be easily sedimented to change solutions. The stripping solutions removed from the zymosan were pooled, their volume was measured, and their absorbance was measured at 510 nm against standards prepared from o-phenanthroline and FeSO₄ (35). This allowed quantitation of the iron originally present in the zymosan: we found zymosan contained approximately 5 nmol of iron per mg. The iron bound to the zymosan was probably Fe³⁺, because solutions without ascorbate liberated no Fe²⁺. After the treatments with ascorbate and o-phenanthroline, the zymosan was washed twice with 10 mM HEPES containing 1 mM DETAPAC, and then boiled for 10 min, as is standard for preparing Zop (30). The zymosan was then opsonized using human serum that had been treated with chelex resin (Bio-Rad) to remove any transition metal ions. Finally, the Zop was washed with 10 mM HEPES and stored at 4°C until it was used. The "control" preparation of Zop was produced by treating a second aliquot of zymosan exactly in parallel, except all chelators and ascorbate were omitted.

Assays of oxidative modification of LDL
LDL oxidation was carried out in 35-mm wells (6-well plates) using 1 million cells per well in serum-free RPMI medium containing 200 µg/ml LDL, in a final volume of 1 ml. In some incubations, Zop was present at a final concentration of 2 mg/ml (25) (26). The incubations, usually in duplicate or triplicate, were performed at 37°C in 95% air/5% CO₂. After 24 h, media were harvested and centrifuged to remove cells and/or Zop. Some experiments used LDL radioiodinated with tyramine cellobiose (20–50 cpm/ng). Small amounts of EDTA that were present in the LDL solution carried over into the incubations (<3 µM); this level of EDTA had no effect on the results, as judged by experiments in which LDL was dialyzed against RPMI prior to incubation with cells (data not shown). Controls for LDL oxidation assays were performed by incubating LDL in RPMI in the absence of cells. Oxidative modification of LDL was routinely assayed by measurement of thiobarbituric acid-reactive substances (TBARS) as previously described (36), measuring fluorescence (excitation at 515 nm, emission at 553 nm).

Other assays
Superoxide production was measured as described (30). Briefly, 1 million cells were incubated in 1 ml of Earles salts containing cytochrome c, in the presence and absence of superoxide dismutase (SOD). After 60 min, the medium was removed, and the SOD-inhibitable reduction of cytochrome c was measured spectrophotometrically. Results are expressed as µM superoxide produced in 1 h. Iron (as Fe²⁺) released into cell culture media was quantitated with bathophenanthroline sulfonate (BPS) as described (37).

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

Characterization of LDL oxidized by Zop-stimulated monocytes
Human peripheral blood monocytes and mouse peritoneal macrophages oxidized LDL in RPMI medium when stimulated by Zop, but not when unstimulated ( Table 1). Our data are similar to data in previous reports (25) (26). There was cell-type specificity to this oxidation because rabbit aortic endothelial cells were unable to oxidize LDL in RPMI whether or not Zop was present (Table 1).

LDL oxidized by activated monocytes was similar to copper-oxidized LDL with respect to a number of biochemical changes that are characteristic of oxidized LDL. In most cases, the characteristics of monocyte-oxidized LDL were intermediate between native LDL and copper-oxidized LDL, indicating that activated monocytes oxidize LDL, but not as heavily as does 5 µM CuSO₄. The characteristics analyzed included migration on agarose gels ( Figure 1), UV absorbance and fluorescence ( Figure 2), and recognition by the macrophage scavenger receptor ( Table 2). Lysophosphatidylcholine content of LDL was also measured, following standard procedures (38) (39). We found that native LDL contained 38 nmol lysophosphatidylcholine per mg LDL protein, and this increased when the LDL was oxidized by 5 µM CuSO₄ (388 nmol/mg) or by Zop-activated monocytes (121 ± 2; mean and range of duplicates). In contrast, lysophosphatidylcholine content of LDL did not significantly increase after incubation of LDL with unactivated monocytes (36 ± 8 nmol/mg) nor with Zop alone in the absence of cells (47 ± 3).

View larger version (39K): [in this window] [in a new window]   Figure 1. Agarose gel electrophoresis of LDL oxidized by stimulated monocytes. 125I-labeled LDL was incubated with monocytes and/or Zop in RPMI under usual conditions (see Materials and Methods), and electronegativity of the samples was compared to native 125I-labeled LDL and copper-oxidized 125I-labeled LDL. Samples were loaded into pre-cast agarose gels and electrophoresed as recommended (Ciba Corning Lipoprotein Electrophoresis System, Palo Alto, CA). The electrophoretogram was subjected to autoradiography. The treatment of the 125I-labeled LDL samples is indicated beneath the lanes; the lane on the far left contained native (unincubated) 125I-labeled LDL. The general pattern of these results was confirmed using Fat Red B staining of non-radioactive LDL preparations that were re-isolated by ultracentrifugal flotation after incubation with monocytes and/or Zop (data not shown).

View larger version (45K): [in this window] [in a new window]   Figure 2. UV and fluorescence spectra of LDL oxidized by stimulated monocytes. LDL was incubated with monocytes and/or Zop in RPMI under usual conditions (see Materials and Methods). The LDL samples were re-isolated by ultracentrifugal flotation, dialyzed, and sterile filtered, and then spectra were recorded at room temperature. A: UV spectra were measured at LDL concentrations of 66 µg/ml as difference spectra versus native (unincubated) LDL. B: Fluorescence emission spectra were measured using LDL samples at 270 µg/ml, with excitation at 350 nm (350 nm was the peak of the excitation spectrum; data not shown). The shaded band indicates the ranges of fluorescence spectra of three different control LDL samples: native LDL, LDL incubated with unstimulated monocytes, and LDL incubated with Zop alone.

Superoxide production does not correlate with LDL oxidation
It has been previously suggested that superoxide causes LDL oxidation (26). Both Zop and tetradecanoyl phorbol acetate (TPA) stimulate monocytes to produce superoxide. We found that the extent and time course of superoxide production was similar for monocytes stimulated with Zop or with TPA ( Figure 3 A). In contrast, LDL oxidation occurred when monocytes were stimulated with Zop, but not when monocytes were stimulated with TPA (Figure 3b). Therefore, superoxide alone did not cause LDL oxidation under these conditions.

View larger version (27K): [in this window] [in a new window]   Figure 3. Superoxide production does not correlate with LDL oxidation. A: Time course of superoxide production by human monocytes incubated with TPA or Zop. The activating agent was added to the monocytes at time zero, and superoxide production was measured at various times. At each time point, superoxide production was measured for 1 h; values represent the production of superoxide over the preceding hour. Unstimulated monocytes produced 1.9 ± 0.5 µM superoxide per hour. B: LDL oxidation by human monocytes incubated with TPA, Zop, or with no activator. Incubations were for 24 hours, as described in Materials and Methods.

Nitric oxide production is not required for LDL oxidation by activated monocytes
It has been reported that nitric oxide can oxidize LDL (40), and that the combination of nitric oxide plus superoxide is a potent oxidant (41). Because of the physiological importance of nitric oxide, we tested whether nitric oxide was required for LDL oxidation, even though human monocytes probably do not produce nitric oxide (42). We measured TBARS production during LDL oxidation by activated monocytes in the presence or absence of N-methyl arginine, which inhibits nitric oxide synthase (43), or dexamethasone, which blocks production of the inducible form of nitric oxide synthase (44). In this experiment, control activated monocytes oxidized LDL and produced 9.9 ± 0.9 nmol/mg TBARS; neither N-methyl arginine (11.7 ± 1.0) nor dexamethasone (8.7 ± 0.2) significantly blocked LDL oxidation. We conclude that nitric oxide production is not required for LDL oxidation by Zop-activated monocytes.

Inhibition of LDL oxidation by various compounds
To further characterize LDL oxidation by activated monocytes, we tested the ability of various compounds to inhibit the oxidation. As shown in Table 3, classic chain-terminating antioxidants such as butylated hydroxytoluene and N,N'-diphenyl-phenylenediamine (5) were effective inhibitors. The chelators EDTA and DETAPAC were found to be potent inhibitors of LDL observation by Zop-activated monocytes. We also found that SOD inhibited LDL oxidation in this system, as previously reported (26).

Zop contains iron, which is necessary for LDL oxidation
The ability of the chelators EDTA and DETAPAC to inhibit LDL oxidation by Zop-activated monocytes implied that transition metal ions are required in this system. The transition metal ions might come from either the Zop or the monocytes. Using o-phenanthroline and ascorbate (see Materials and Methods), we found that zymosan contains approximately 5 nmol of Fe per mg, probably as Fe³⁺. This result was confirmed by inductively coupled plasma atomic emission spectroscopy, which gave a value of 4.2 nmol of Fe per mg of zymosan (analysis done at the Soil Testing Laboratories, University of Minnesota, St. Paul, MN). Copper was also present at 0.14 nmol per mg. The relative significance of the iron and copper is discussed below.

To determine whether the iron associated with the zymosan was important for the biochemical activity of zymosan, we compared monocytes treated with Zop to monocytes treated with Zop that had been stripped of its iron. Zymosan was stripped of iron using ascorbate plus o -phenanthroline (see Materials and Methods). The iron-stripped Zop supported superoxide production by monocytes and macrophages, but did not support LDL oxidation ( Table 4).

Monocytes cause release of Fe²⁺ from Zop
We hypothesized that iron present in the Zop preparations was released as Fe²⁺ during incubation with cells, and that the Fe²⁺ then participated in LDL oxidation. To quantitate release of Fe²⁺ from Zop, cells were incubated in RPMI with Zop stripped of its iron or with control Zop, in the presence and absence of the Fe²⁺ reagent bathophenanthroline sulfonate (BPS). After a 24 h incubation, the media were harvested, centrifuged, and the iron–BPS complex was quantitated. As shown in Table 5, both HL-60 cells and human peripheral blood monocytes cause the release of iron from Zop but not from Zop that had been stripped of iron. Based on the iron content of zymosan measured by atomic emission spectroscopy, the cells caused the release of approximately one-third of the total zymosan-associated iron (2.7 µM iron in 1 ml conditioned medium divided by 8.4 nmol iron present in 2 mg zymosan in the incubation = 32%). Our assay detected no iron when monocytes or Zop were incubated in the absence of the other (data not shown). Comparison of Table 4 and Table 5 suggest that iron released from Zop is required for LDL oxidation by activated monocytes.

Replenishing the iron allows stripped Zop to support LDL oxidation
To determine whether iron was the critical factor lost from zymosan during stripping, iron was added back to incubations with stripped Zop. Reagent FeSO₄ was added to achieve the same amount that was found to be released from the commercial zymosan by the cells. Reagent CuSO₄ was also tested, because the presence of 0.14 nmol copper per mg of zymosan had been detected by inductively coupled plasma atomic emission spectroscopy. In the experiment, HL-60 cells incubated with iron-stripped Zop and LDL produced 0.8 ± 0.1 nmol/mg of TBARS. The addition of 0.15 µM CuSO₄ did not increase the production of TBARS (0.9 ± 0.1), whereas the addition of 3 µM FeSO₄ significantly increased TBARS production to 9.5 ± 0.2 nmol/mg. The addition of 3 µM FeSO₄ to RPMI medium did not enhance LDL oxidation in the absence of cells. This last observation is similar to data published by Lamb and Leake (18). Taken together, these data imply that the iron present in the zymosan is important for LDL oxidation, and also that the procedure used to strip the zymosan did not introduce an antioxidant that prevents LDL oxidation by monocytes.

Contact between Zop and the cells is not required for LDL oxidation
To test whether contact between Zop and the cells was required for iron release we used Trans-well inserts. These inserts fit inside the wells of 6-well plates, and are made of a microporous membrane with 0.25-micron pores such that soluble molecules pass through freely, but particulates are held in place approximately 1 mm above the cells. When HL-60 cells were placed in the wells and Zop was placed in the insert such that there was no contact, very little LDL oxidation occurred ( Table 6). When TPA was also added, however, there was significant LDL oxidation, even though cells plus TPA alone caused no LDL oxidation (Table 6). This suggests that activated monocytes secrete a factor that causes iron release from Zop, leading to LDL oxidation. This factor is released by cells stimulated by the Zop itself or by TPA.

We also performed a control experiment to show that there was no leakage of the Zop from the insert onto the cells. Zop is particulate and is prepared via repeated low-speed sedimentations (30), so it would be expected that the membrane pores would retain the Zop in the insert. To confirm this, we measured superoxide production by HL-60 cells incubated with empty inserts or with inserts containing Zop. HL-60 cells alone produced 1.6 ± 0.4 µM superoxide per h, and with Zop in the inserts, the cells produced 1.8 ± 0.3 µM/h (mean ± SEM of triplicate incubations). Superoxide production increased to 15.8 ± 2.1 µM/hr when the Zop was placed directly onto the cells in the standard method. This experiment showed that Zop in the insert does not leak through to the cells.

Zop alone, but not iron-stripped Zop, can oxidize LDL
We noticed that control incubations of LDL with Zop in the absence of cells gave slightly higher TBARS than LDL incubated in medium alone. We therefore tested whether Zop alone can oxidize LDL over many days at 37°C. LDL was incubated in the absence of cells at 37°C in RPMI medium with or without Zop or iron-stripped Zop. After 48 h, TBARS (nmol/mg) produced were 0.3 ± 0.03 without Zop, 0.5 ± 0.05 with iron-stripped Zop, and 7.0 ± 0.2 with Zop. The TBARS increased after 96 h to 0.5 ± 0.04 , 0.8 ± 0.07, and 11 ± 0.1 , respectively. These data suggest that the iron carried by Zop is capable of causing limited LDL oxidation in the absence of cells.

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

LDL oxidation by cultured cells has been intensively studied (1) (11) (12) (13) (14) (15) (16) (17) (18) (19) (20) (21) (22) (23) (24) (25) (26) (27) (28) (29). Transition metal ions are usually required for this process (14) (15) (16) (17) (18) (19) (20) (21) (22) (23) (24). Free transition metal ions may not be available in vivo, however, due to the presence of endogenous chelators. Thus it would be of great interest if there existed an in vitro cellular mechanism for LDL oxidation that was independent of transition metal ions. Activated monocytes are known to secrete oxidizing agents (30), and it was reported that monocytes activated by Zop could oxidize LDL in RPMI medium, which is formulated without transition metal ions (25) (26). This was an intriguing candidate for a possible physiologically relevant mechanism for cellular LDL oxidation. We have further characterized this system and found that the oxidized LDL produced is similar to copper-oxidized LDL. The monocyte-oxidized LDL contained lysophosphatidylcholine, had increased mobility on agarose gels, displayed increased fluoresence, and was a ligand for scavenger receptors. In general, the absolute extent of LDL oxidation by activated monocytes was less than the oxidation achieved by copper ions.

We have also addressed the mechanism of LDL oxidation by activated monocytes. We found that, despite the absence of transition metal ions in the formulation for RPMI medium, the LDL oxidation required transition metal ions, as judged by the fact that chelators inhibited the oxidation. We discovered that small amounts of iron were present in the commercial zymosan used to prepare the Zop. The ability of Zop to stimulate LDL oxidation was completely lost when the Zop was subjected to a procedure that stripped it of iron, even though iron-stripped Zop still activated the cells. Commercial zymosan preparations apparently are contaminated with sufficient iron to support LDL oxidation by activated monocytes. Thus activated monocytes require transition metal ions to oxidize LDL, as do other cell types (14) (15) (16) (17) (18) (19) (20) (21) (22) (23) (24).

If Zop carries iron, why doesn't Zop alone cause LDL oxidation? Although Zop alone does catalyze minimal LDL oxidation, Zop plus monocytes oxidize LDL to a much greater extent. Our data suggest that activated monocytes secrete a factor that reduces Zop-bound Fe³⁺ to Fe²⁺, and the Fe²⁺ is released from the Zop and participates in LDL oxidation. This factor is likely to be superoxide, given the known ability of superoxide to reduce Fe³⁺ in ferritin, followed by iron-catalyzed lipid peroxidation (37) (45). This action of superoxide would explain the ability of SOD to inhibit LDL oxidation (Table 3 and reference 26). In our experiments, superoxide alone did not directly oxidize LDL, as judged by the fact that stimulation of monocytes with TPA, which caused substantial superoxide production, did not cause LDL oxidation. Stimulation of monocytes with Zop, however, caused superoxide production and also caused LDL oxidation. The ability of Zop to stimulate LDL oxidation was completely lost when the Zop was subjected to a procedure that stripped it of iron, even though iron-stripped Zop still supported superoxide production by the cells. Thus we suggest that the superoxide, or some other factor released by activated monocytes, liberates contaminating iron in the commercial zymosan preparations, which is followed by iron-catalyzed LDL oxidation. The liberated iron may interact with superoxide molecules to cause LDL oxidation, e.g., via Haber-Weiss reactions. Hiramatsu et al. (46) have previously shown that cellular superoxide production can cause LDL oxidation in the presence of iron. It is also worth noting that Garner et al. (47) have shown that macrophages can reduce transition metal ions by superoxide-independent mechanisms.

There is disagreement in the literature as to whether superoxide alone can cause LDL oxidation. Production of superoxide by radiolysis can lead to LDL oxidation, although this oxidation is efficient only at pH below physiological levels (48). Lynch and Frei (49) used xanthine plus xanthine oxidase to produce superoxide and showed that superoxide only oxidizes LDL in the presence of transition metal ions. The elegant work of Hiramatsu et al. showed (46) that TPA-stimulated human monocytes could oxidize LDL only if they could produce superoxide. A critical observation in this study was that 10 µM EDTA inhibited 85% of the LDL oxidation catalyzed by activated monocytes (46), implying that transition metal ions were required. Transition metal ions, especially iron, are common contaminants in commercial reagents and even in distilled water stored in glass; their presence may, however, vary among different laboratories. To summarize, it is clear that superoxide plus transition metal ions can cause LDL oxidation and lipid peroxidation in general ((21), (37), (45), (46), (49) (50) (51) and the present work). It is unlikely that superoxide alone is capable of oxidizing LDL at physiological pH.

Which of the many proposed mechanisms for LDL oxidation are most likely to be relevant in vivo? In the presence of transition metal ions, cells can cause LDL oxidation by secretion of superoxide, as discussed above. Superoxide production occurs in vivo, and superoxide can spontaneously dismutate to H₂O₂. These molecules are oxidants, but as discussed above, their ability to significantly oxidize LDL appears to be dependent on transition metal ions. Free transition metal ions may or may not be available in vivo. Total body iron was shown to be correlated with carotid artery atherosclerosis in humans (52), but iron overload did not increase atherosclerosis in hypercholesterolemic rabbits (53). Atherosclerotic "gruel" has been reported to contain reactive iron that is competent for lipid peroxidation (54). LDL oxidation has been achieved in the absence of transition metal ions by enzymatic systems using peroxidase (55), lipoxygenase (36) (56), or myeloperoxidase (57). Atherosclerotic lesions have been shown to contain both lipoxygenase (58) and myeloperoxidase (59). Future research may clarify the important issue of whether LDL oxidation in vivo requires free transition metal ions, or if enzymes present in the lesion can oxidize LDL without free transition metal ions.

	FOOTNOTES

¹ Current address: Immunomedics Inc., 300 American Road, Morris Plains, NJ 07950.

	ACKNOWLEDGMENTS

The authors wish to thank Dr. Hugh Rosen for many helpful discussions about activated monocytes. We thank Dr. Steve Grant for advice on cellular nitric oxide production. The authors are grateful to Dr. Yu-sheng Chao for his support and advice.

Manuscript received April 10, 1998; and in revised form July 8, 1998.

Abbreviations: BPS, bathophenanthroline sulfonate; DETAPAC, diethylenetriaminepentaacetic acid; SOD, superoxide dismutase; TBARS, thiobarbituric acid-reactive substances; TPA, tetradecanoyl phorbol acetate; Zop, opsonized zymosan; apo, apolipoprotein; LDL, low density lipoprotein(s)

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TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

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