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Journal of Lipid Research, Vol. 48, 565-582, March 2007
Copyright © 2007 by American Society for Biochemistry and Molecular Biology

Import and fate of fluorescent analogs of oxidized phospholipids in vascular smooth muscle cells

Alexandra Moumtzi^1,*, Michael Trenker^1,*, Karlheinz Flicker^*, Elfriede Zenzmaier^*, Robert Saf and Albin Hermetter^2,*

^* Institute of Biochemistry, Graz University of Technology, A-8010 Graz, Austria
Institute for Chemistry and Technology of Organic Materials, Graz University of Technology, A-8010 Graz, Austria

Published, JLR Papers in Press, November 29, 2006.

¹ A. Moumtzi and M. Trenker contributed equally to this work.

² To whom correspondence should be addressed. e-mail: albin.hermetter{at}tugraz.at

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Lipid oxidation is now thought to be an initiating and sustaining event in atherogenesis. Oxidatively fragmented phospholipids, namely 1-palmitoyl-2-glutaroyl-sn-glycero-3-phosphocholine (PGPC) and 1-palmitoyl-2-(5-oxovaleroyl)-sn-glycero-3-phosphocholine (POVPC), present in minimally modified LDL and atherosclerotic lesions, have been reported to elicit a wide range of pathophysiological responses in the cells of the vascular wall. Nevertheless, the question of their potential sites of action and their primary molecular targets remains open. To address this issue, a series of fluorescently labeled analogs, which differ with regard to structure and binding site of the fluorophore, were synthesized and used as tools for studying the uptake, intracellular stability, and distribution of PGPC and POVPC in vascular smooth muscle cells (VSMCs). We demonstrate that in accordance with their lysophospholipid-like structure, these highly similar molecules transferred rapidly either from aqueous phospholipid dispersions or preloaded native LDL into VSMCs, producing disparate fluorescence patterns irrespective of the attached fluorophore. PGPC derivatives were translocated to the lysosomes. In sharp contrast, POVPC analogs were initially captured in the plasma membrane, most likely in consequence of the formation of covalent adducts with free amino and sulfhydryl groups of proteins and phospholipids. LDL internalization is not required for cellular lipid uptake. Collectively, our data provide evidence that oxidized phospholipids, owing to their high exchangeability between lipoproteins and cell membranes, may act within a short time on different cellular sites in VSMCs and affect various lipid and protein components through physical or chemical interactions, which might then serve as starting points for intracellular signaling.

Supplementary key words atherosclerosis • lipoproteins • lipid synthesis • lipid transport

Abbreviations: AcOH, acetic acid; Alexa647, Alexa Fluor 647; apoB, apolipoprotein B-100; BODIPY, 4,4-difluoro-4-bora-3a,4a-diaza-s-indacene; 2-D, two-dimensional; DCC, N,N'-dicyclohexylcarbodiimide; DMAP, p-(N,N-dimethylamino)pyridine; ER, endoplasmic reticulum; EtOH, ethanol; FCS, fetal calf serum; HNE, 4-hydroxy-2-nonenal; MALDI-TOF, matrix-assisted laser desorption/ionization time-of-flight; MeOH, methanol; PGPC, 1-palmitoyl-2-glutaroyl-sn-glycero-3-phosphocholine; PGPE, 1-palmitoyl-2-glutaroyl-sn-glycero-3-phosphoethanolamine; PLA₂, phospholipase A₂; PLD, phospholipase D; POPE, 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine; POVPC, 1-palmitoyl-2-(5-oxovaleroyl)-sn-glycero-3-phosphocholine; POVPE, 1-palmitoyl-2-(5-oxovaleroyl)-sn-glycero-3-phosphoethanolamine; PyrGPC, 1-(10-pyrenedecanoyl)-2-glutaroyl-sn-glycero-3-phosphocholine; PyrOPC, 1-(10-pyrenedecanoyl)-2-oleoyl-sn-glycero-3-phosphocholine; PyrOVPC, 1-(10-pyrenedecanoyl)-2-(5-oxovaleroyl)-sn-glycero-3-phosphocholine; R_f, relative mobility; SE, succinimidyl ester; VSMC, vascular smooth muscle cell

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Oxidative modification of LDL is considered an important pathogenetic factor in atherosclerosis (1–3). Preferential targets of oxidative attack are the esterified polyunsaturated fatty acids in the phospholipids of biological membranes and the shell that surrounds the insoluble lipids of lipoproteins (4). Oxidative fragmentation of arachidonoyl phospholipids results in shortened sn-2 residues with or without -aldehydic, -hydroxy, or -carboxy functions (5, 6). Such compounds are also formed in vitro by oxidation of 1-palmitoyl-2-arachidonoyl-sn-glycero-3-phosphocholine (7, 8). It has been demonstrated that the truncated sn-2 substituents of these compounds are major determinants of their bioactivity (9). Studies from several laboratories have revealed that the biological effects triggered by minimally modified LDL can largely be attributed to phospholipid oxidation products (7, 8, 10, 11). Their increased levels in atherosclerotic plaques (12, 13) and the increased antibody titers against oxidized phospholipids in humans and mice with lesions (14–16) attract attention to the pathological relevance of these molecules.

A rapidly growing interest has been focused on two major representatives in the series of homologous oxidized phospholipids, namely 1-palmitoyl-2-glutaroyl-sn-glycero-3-phosphocholine (PGPC) and 1-palmitoyl-2-(5-oxovaleroyl)-sn-glycero-3-phosphocholine (POVPC). Their importance is stressed by the finding that they selectively activate processes in vascular cells that may contribute to the pathogenesis of atherosclerosis as well as other chronic inflammatory diseases. It is noteworthy that, in spite of their high structural similarity, due to a single hydrophobic chain, a bulky polar head group, and the polar sn-2 acyl fragment, they partially exert different cellular effects. PGPC and POVPC exhibit proinflammatory actions (7, 17–19) but also increase the synthesis of protective enzymes such as heme oxygenase 1 (20). In human vascular smooth muscle cells (VSMCs), POVPC and, to a lesser extent, PGPC activated apoptotic signaling pathways via the stimulation of acid sphingomyelinase (11).

VSMCs can acquire a broad spectrum of different phenotypes in response to changes in a variety of physiological or pathological factors (21, 22), including oxidized phospholipids (11, 18, 23). Nonetheless, the primary molecular targets and the mechanisms of their activation by these molecules are largely unknown. The identification of the cell compartments that accumulate oxidized phospholipids is of great importance in defining the mode of action by which they cause cellular dysfunction.

This study was designed to gain insight into the import, intracellular distribution, and potential targets of PGPC and POVPC in VSMCs. For this purpose, we synthesized different fluorescent analogs of the oxidized phospholipids. Using fluorescence microscopy, we found rapid uptake of both phospholipid species but substantial differences with regard to their intracellular accumulation. PGPC derivatives were initially transferred to perinuclear regions, corresponding to lysosomes and partially to the endoplasmic reticulum (ER), whereas labeled POVPC became highly enriched in the plasma membrane at early time points. These data support the assumption that multiple protein and lipid interaction sites must exist in VSMCs that are affected soon after their initial contact with oxidized phospholipids, thus initiating intracellular signaling.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Materials
Chemicals for lipid synthesis, N,N'-dicyclohexylcarbodiimide (DCC; purity > 98%), p-(N,N-dimethylamino)pyridine (DMAP; 99%), phospholipase A₂ (PLA₂) from Naja naja venom in Tris buffer (pH 8.9) (EC 3.1.1.4; specific activity, 1,000 IU/mg), and phospholipase D (PLD) type VII from Streptomyces species in Tris buffer (pH 5.6) (EC 3.1.4.4; specific activity, 500 IU/mg) were obtained from Sigma-Aldrich. 4,4-Difluoro-4-bora-3a,4a-diaza-s-indacene-succinimidyl ester (BODIPY^TMSE), Alexa^TM Fluor 647 (Alexa647)-SE, and 1-pyrenedecanoic acid were purchased from Molecular Probes (Eugene, OR). Chemicals for gel electrophoresis were from Bio-Rad Laboratories (Hercules, CA), unless noted otherwise. The dye reagent for the Bradford protein assay was also purchased from Bio-Rad. Organic solvents and all other reagents were obtained from Merck (Darmstadt, Germany), unless indicated otherwise. Tissue culture dishes and flasks were obtained from Sarstedt (Nürmbrecht, Germany). Media and supplements for cell culture were purchased from PAA Laboratories (Linz, Austria). Ninety-six-well microtiter plates for protein determinations were obtained from Greiner (Kremsmünster, Austria).

Cell culture
Rat aortic smooth muscle cells (A7r5; catalog No. CRL-1444, smooth muscle, thoracic aorta, DBIX rat; American Type Culture Collection, Rockville, MD) were cultivated in DMEM supplemented with 10% fetal calf serum (FCS), 2 mM L-glutamine, 100 U/ml penicillin/streptomycin, and 25 mM HEPES at 37°C in a humidified CO₂ (8%) atmosphere. Incubations with fluorescent phospholipids were routinely carried out in serum-free culture medium at 37°C in a CO₂ incubator after washing cells twice with PBS to remove residual FCS. Working solutions of the labeled lipids were prepared by diluting dispersions of these compounds in PBS with the appropriate amount of incubation medium.

Synthesis of aliphatic and fluorescent phospholipids
TLC was routinely performed on aluminum plates precoated with 0.25 mm of silica gel 60 (Merck, Darmstadt, Germany). CHCl₃/methanol (MeOH)/water (65:25:4, v/v/v) was used as the developing system, unless indicated otherwise. Spots were visualized by illumination under short-wavelength ultraviolet light (fluorophores) or by treatment with molybdenum blue reagent (phosphorus) (24), 50% H₂SO₄ (organic material), or ninhydrin (amino groups). Anhydrous dichloromethane was obtained by distillation over phosphorus pentoxide. Flash column chromatography was performed on 230–400 mesh silica gel supplied by Merck. Fluorescence spectroscopic measurements of methanolic solutions of the fluorescent phospholipids were conducted in quartz cuvettes using a spectrofluorophotometer (RF-3501 PC; Shimadzu, Korneuburg, Austria). The concentrations and quantities of lipids were determined by phosphorus analysis (25) or by absorption measurements of the fluorescent compounds.

Synthesis of PGPC
PGPC (Fig. 1A , compound 1) was synthesized according to a modified version of the procedure of Watson et al. (12) and Subbanagounder, Watson, and Berliner (26). A solution of dry 1-palmitoyl-sn-glycero-3-phosphocholine (62 mg, 125 µmol), dry glutaric anhydride (70 mg, 613 µmol, 5 eq), and DMAP (75 mg, 614 µmol, 5 eq) in 6 ml of anhydrous dichloromethane was magnetically stirred overnight at 35°C. The reaction was monitored by TLC with CHCl₃/MeOH/25% NH₃ (65:35:5, v/v/v) as a developing system and quenched by the addition of 3 ml of MeOH. The resulting mixture was washed once with 1.8 ml of MeOH/water (1:1, v/v). After evaporation of the organic phase, excess DMAP was removed by trituration with 4 ml of diethylether. Removal of the supernatant gave 32 mg of PGPC [42%; relative mobility (R_f) = 0.05].

View larger version (12K): [in this window] [in a new window]   Fig. 1. Syntheses of 4,4-difluoro-4-bora-3a,4a-diaza-s-indacene (BODIPY) and pyrene analogs of 1-palmitoyl-2-glutaroyl-sn-glycero-3-phosphocholine (PGPC) and 1-palmitoyl-2-(5-oxovaleroyl)-sn-glycero-3-phosphocholine (BODIPY-POVPC). A: BODIPY-1-palmitoyl-2-glutaroyl-sn-glycero-3-phosphoethanolamine (BODIPY-PGPE). B: BODIPY-1-palmitoyl-2-(5-oxovaleroyl)-sn-glycero-3-phosphoethanolamine (POVPE). C: 1-(10-Pyrenedecanoyl)-2-glutaroyl-sn-glycero-3-phosphocholine (PyrGPC) and 1-(10-pyrenedecanoyl)-2-(5-oxovaleroyl)-sn-glycero-3-phosphocholine (PyrOVPC). DCC, N,N'-dicyclohexylcarbodiimide; DMAP, p-(N,N-dimethylamino)pyridine; PLA2, phospholipase A2; PLD, phospholipase D; SE, succinimidyl ester.

Synthesis of BODIPY-PGPE
To a magnetically stirred solution of BODIPY-SE (0.96 mg, 2.5 µmol) in 1 ml of CHCl₃/MeOH (2:1, v/v) were added PGPE (4.4 mg, 7.8 µmol, 3 eq) (Fig. 1A, compound 2) and triethylamine pro analysi (p.a.) (10 µl, 72 µmol, 30 eq) (Fig. 1A, step 2). Then the flask was flushed with nitrogen and protected from light, and the resulting solution was stirred at room temperature for 1 h. The progress of the reaction was monitored by TLC. The solvent was removed under a nitrogen stream until a volume of 400 µl was reached, from which the lipid was purified by preparative TLC. The desired compound was visualized under ultraviolet light, scraped off the TLC plate, and eluted three times from the silica gel with CHCl₃/MeOH (1:4, v/v). The solvent was removed from the combined extracts by rotary evaporation to give BODIPY-PGPE (1.77 mg, 85%; R_f = 0.28) (Fig. 1A, compound 3).

Synthesis of Alexa647-PGPE
Alexa647-PGPE corresponds to BODIPY-PGPE (Fig. 1A, compound 3) but contains Alexa647 instead of BODIPY on its head group. It was obtained from Alexa647-SE (0.5 mg, 0.4 µmol) and PGPE (0.68 mg, 1.2 µmol, 3 eq) (Fig. 1A, compound 2) in 0.5 ml of CHCl₃/MeOH (2:1, v/v) after the addition of triethylamine p.a. (5 µl, 36 µmol, 90 eq) followed by stirring at room temperature for 90 min. The reaction mixture was strictly protected from light. The progress of the reaction was monitored by TLC (RP-18 F_254s; Merck). The product was purified by preparative TLC with water/ethanol (EtOH)/n-propanol (20:57:23, v/v/v) as a developing system, scraped off, and eluted twice with CHCl₃/MeOH (1:1, v/v) and once with MeOH. Removal of the solvent under a stream of nitrogen gave Alexa647-PGPE (0.26 µmol, 65%; R_f = 0.80).

Synthesis of 1-palmitoyl-2-(5-dimethoxypentanoyl)-sn-glycero-3-phosphoethanolamine
A solution of 5,5-dimethoxypentanoic acid methyl ester (220 mg, 1.25 mmol) (Fig. 1B, compound 4) and sodium hydroxide (250 mg, 6.25 mmol, 5 eq) in 5 ml of water/MeOH/tetrahydrofuran (2:5:3, v/v/v) was stirred at room temperature for 90 min (Fig. 1B, step 1). After cooling to 0°C, the reaction mixture was acidified to pH 2.1 by subsequent addition of 6 ml of 1 N HCl and appropriate amounts of 0.1 N HCl and then extracted with dichloromethane (3 x 15 ml). The combined organic extracts were washed with water (2 x 10 ml) and dried over Na₂SO₄. The solvent was removed under reduced pressure except for 10–15 ml containing the desired product (R_f = 0.35) (Fig. 1B, compound 5), which was immediately used for the acylation reaction without further purification. 1-Palmitoyl-sn-glycero-3-phosphocholine (209 mg, 0.42 mmol) (Fig. 1B, compound 6), DCC (270 mg, 1.3 mmol, 3 eq), and DMAP (160 mg, 1.3 mmol, 3 eq) were added to this solution of 5,5-dimethoxypentanoic acid (Fig. 1B, step 2). The mixture was stirred under nitrogen at room temperature overnight. The reaction was monitored by TLC. After the addition of 6 ml of MeOH, the organic solution was washed twice with MeOH/water (1:1, v/v). The solvent was removed under vacuum, and traces of water were evaporated after the addition of benzene/EtOH (3:2, v/v). The oily residue was flash chromatographed on 12 g of silica gel with CHCl₃/MeOH/25% NH₃ (65:35:5, v/v/v) as a solvent to give 1-palmitoyl-2-(5,5-dimethoxypentanoyl)-sn-glycero-3-phosphocholine (245 mg, 91%; R_f = 0.14) (Fig. 1B, compound 7). The ethanolamine analog (Fig. 1B, compound 8) was obtained by transphosphatidylation of the choline lipid as described previously for the conversion of PGPC to PGPE (compare Fig. 1A, step 1) [yield, 6.2 mg, 66%; R_f = 0.24 using CHCl₃/MeOH/25% NH₃ (65:35:5, v/v/v) as a developing solvent] (Fig. 1B, step 3).

Synthesis of 1-palmitoyl-2-(5-oxovaleroyl)-sn-glycero-3-phospho-N-(3-BODIPY-propionyl)-ethanolamine
Triethylamine p.a. (10 µl, 72 µmol, 14 eq) was added to a solution of 1-palmitoyl-2-(5,5-dimethoxypentanoyl)-sn-glycero-3-phosphoethanolamine (3.0 mg, 5.0 µmol) (Fig. 1B, compound 8) and BODIPY-SE (1.95 mg, 5.0 µmol, 1 eq) in 1 ml of CHCl₃/MeOH (2:1, v/v) (Fig. 1B, step 4). The reaction mixture was stirred at room temperature for 80 min. The solvent was removed under vacuum, yielding 1-palmitoyl-2-(5,5-dimethoxypentanoyl)-sn-glycero-3-phospho-N-(3-BODIPY-propionyl)-ethanolamine (Fig. 1B, compound 9), which was suitable for the next reaction without further purification. Release of the desired product was accomplished by acetal cleavage of this stable precursor (2.0 mg, 2.3 µmol) with 400 µl of tetrahydrofuran-HCl (1 N) (Fig. 1B, step 5). After only 2 min, the reaction mixture was neutralized with NaHCO₃ followed by extraction of the product with 1.2 ml of CHCl₃/MeOH (2:1, v/v). The organic phase was washed twice with 300 µl of water, dried over Na₂SO₄, and evaporated under reduced pressure, leading to 1.65 mg of BODIPY-1-palmitoyl-2-(5-oxovaleroyl)-sn-glycero-3-phosphoethanolamine (POVPE) [87%; R_f = 0.22 using CHCl₃/MeOH/water (15:5:0.1, v/v/v) as a developing system] (Fig. 1B, compound 10).

Synthesis of 1-palmitoyl-2-oleoyl-sn-glycero-3-phospho-N-(3-BODIPY-propionyl)-ethanolamine
BODIPY-SE (1.00 mg, 2.57 µmol), triethylamine p.a. (10 µl, 72 µmol, 28 eq), and 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine (POPE; 5.53 mg, 7.70 µmol, 3 eq) were dissolved in 1 ml of CHCl₃/MeOH (2:1, v/v). The mixture was stirred at room temperature for 60 min, and the solvent was removed under a stream of nitrogen. The white oily residue was dissolved in 500 µl of CHCl₃/MeOH (2:1, v/v). The product was purified by preparative TLC. The fluorescent band containing the product was scraped off the plate and eluted three times with CHCl₃/MeOH (1:4, v/v). The combined extracts were evaporated to deliver the desired product (1.31 mg, 51%; R_f = 0.59).

Synthesis of 1-(10-pyrenedecanoyl)-2-glutaroyl-sn-glycero-3-phosphocholine
To a magnetically stirred emulsion of 1,2-bis(10-pyrenedecanoyl)-sn-glycero-3-phosphocholine (45 mg, 47 µmol) (Fig. 1C, compound 11) in a mixture of 3 ml of 0.1 M Tris-HCl buffer (pH 8) containing 0.1 M CaCl₂ and 3 ml of diethylether (peroxide-free) were added 50 units of PLA₂ (Naja naja venom) (Fig. 1C, step 1). The reaction mixture was stirred overnight at 35–40°C. After removing the diethylether, the product was extracted from the aqueous solution with CHCl₃/MeOH (2:1, v/v) (3 x 5 ml). The combined organic fractions were evaporated and the residual water was removed under high vacuum, leading to 50 mg of a mixture of 1-(10-pyrenedecanoyl)-sn-glycero-3-phosphocholine (Fig. 1C, compound 12) and free pyrenedecanoic acid. The latter compound was removed by trituration with diethylether. After removal of the solvent under vacuum, 25 mg of the pure lysophospholipid was obtained [88%; R_f = 0.12 in CHCl₃/MeOH/AcOH/water (50:30:10:5, v/v/v/v) as a developing system]. Glutaric anhydride (12 mg, 105 µmol, 11 eq) and anhydrous DMAP (4 mg, 33 µmol, 3 eq) were added to a solution of the lysophospholipid (6.0 mg, 9.8 µmol) in 3 ml of anhydrous dichloromethane (Fig. 1C, step 2a). The reaction was stirred overnight at 35–40°C. Flash chromatography of the crude product on 10 g of silica gel with CHCl₃/MeOH/water (65:25:4, v/v/v) led to 1-(10-pyrenedecanoyl)-2-glutaroyl-sn-glycero-3-phosphocholine (PyrGPC) [2.2 mg, 31%; R_f = 0.18 in CHCl₃/MeOH/AcOH/water (50:30:10:5, v/v/v/v) as a developing system] (Fig. 1C, compound 14).

Synthesis of 1-(10-pyrenedecanoyl)-2-(5-oxovaleroyl)-sn-glycero-3-phosphocholine
1-(10-Pyrenedecanoyl)-sn-glycero-3-phosphocholine (10 mg, 16 µmol) (Fig. 1C, compound 12) was acylated with compound 5 in 10 ml of dichloromethane containing DCC (200 mg, 1.0 mmol, 59 eq) and DMAP (200 mg, 1.6 mmol, 100 eq). The reaction mixture was stirred at room temperature overnight (Fig. 1C, step 2b). The progress of the reaction was monitored by TLC until the reaction was stopped by the addition of 6 ml of MeOH. The resultant solution was washed twice with MeOH/water (1:1, v/v), and the solvent was removed under reduced pressure. The crude product was purified by preparative TLC, scraped off, and eluted five times with 4 ml of CHCl₃/MeOH (1:4, v/v). Residual silica gel was removed by washing the combined organic fractions (concentrated to a volume of 5 ml) with 1 ml of MeOH/water (1:1, v/v). Evaporation of the solvent yielded 6 mg of 1-(10-pyrenedecanoyl)-2-(5,5-dimethoxypentanoyl)-sn-glycero-3-phosphocholine (49%; R_f = 0.17) (Fig. 1C, compound 15). Deprotection of compound 15 (5.6 mg, 7.4 µmol) was performed in analogy to the acid-catalyzed hydrolysis of compound 9 (compare Fig. 1B, step 5), producing 1.1 mg of 1-(10-pyrenedecanoyl)-2-(5-oxovaleroyl)-sn-glycero-3-phosphocholine (PyrOVPC) (21%; R_f = 0.05) (Fig. 1C, step 3).

Synthesis of 1-(10-pyrenedecanoyl)-2-oleoyl-sn-glycero-3-phosphocholine
Oleic acid (99%) (10 µl, 28 µmol, 3 eq), anhydrous DCC (15 mg, 73 µmol, 9 eq), and DMAP (15 mg, 123 µmol, 15 eq) were added to a solution of compound 12 (5.0 mg, 8.2 µmol) in 3 ml of anhydrous dichloromethane. The mixture was stirred at room temperature overnight. The product was purified by preparative TLC. The fluorescent band containing the product was scraped off the plate and eluted three times with CHCl₃/MeOH (1:4, v/v). The combined extracts were evaporated to deliver pure 1-(10-pyrenedecanoyl)-2-oleoyl-sn-glycero-3-phosphocholine (PyrOPC) (3.68 mg, 51%; R_f = 0.13).

Mass spectrometry
Matrix-assisted laser desorption/ionization time-of-flight (MALDI-TOF) mass spectrometry was performed on a Micromass TofSpec2E apparatus equipped with a nitrogen laser ( = 337 nm, operated at 5 Hz) and a time-lag focusing unit. Ions were generated by irradiation just above the threshold laser power. The spectra were recorded in the reflectron mode with an acceleration voltage of 20 kV and externally calibrated with a suitable mixture of poly(ethylene glycol)s. Samples were typically prepared by mixing solutions of the matrix (2,5-dihydroxybenzoic acid; concentration = 10 mg/ml, CH₃CN/0.1% trifluoroacetic acid, 70:30, v/v), the analyte (c = 0.01–1 mg/ml, CHCl₃/MeOH, 2:1, v/v), and Na-trifluoroacetic acid (c = 1 mg/ml, CH₃CN/water, 70:30, v/v) in a ratio of 10:1:0.5 (v/v/v). A 0.5 µl aliquot of the mixture was deposited on the sample plate (stainless steel) and allowed to dry under air. The spectra of 50–100 shots were averaged to improve the signal-to-noise ratio. All m/z values discussed in this work correspond to the most intense peak of any isotope distribution.

Isolation, labeling, and enrichment of LDL with fluorescent phospholipids
Human LDL (1.019–1.069 g/ml) was isolated from pooled fresh plasma of normal individuals by density ultracentrifugation as described (27), filter-sterilized, and stored at 4°C until use within 1 week of isolation. Lipoprotein concentration is expressed in terms of protein content, which was measured by the method of Lowry et al. (28). Lipoprotein labeling with BODIPY-SE was performed as reported previously by Hofer et al. (29). For LDL loading with fluorescent phospholipids, 750 nmol of PyrGPC or 1.25 µmol of PyrOVPC was solubilized in CHCl₃/MeOH (2:1, v/v) and transferred into a pyrex tube. After removing the solvent under a stream of nitrogen, the dry lipid film was overlaid with 250 µg of LDL protein in 190 µl of PBS, dispersed by gentle agitation, and incubated for 30 min at room temperature under nitrogen in the dark. The resulting solution was adjusted to 5 ml with serum-free cell culture medium, leading to a final concentration of 50 µg protein/ml for LDL, 3 nmol/µg protein for PyrGPC, and 5 nmol/µg protein for PyrOVPC.

Fluorescence microscopy
For microscopy experiments, monolayer cultures of A7r5 cells were grown to low confluence in 60 mm Petri dishes containing three coverslips each. Phospholipid concentrations in serum-free culture medium depended on fluorophore emission intensities, individual uptake rates, and the capabilities of the technical equipment.

Import of lipids into cells Experiments were performed with concentrations ranging from 0.1 µM for the BODIPY analogs and Alexa647-PGPE, to 0.5 µM particularly for BODIPY-POVPE, and to 2.0 µM for all pyrene derivatives. Incubation times were: 5, 15, 30, 45, 60, and 90 min (37°C). After treatment, cells were carefully rinsed with PBS and observed with an Axiovert 35 inverted fluorescence microscope equipped with a 100 x 1.3 numerical aperture Plan-Neofluar objective (Carl Zeiss, Oberkochen, Germany), a mercury-arc lamp (HBO® 50W/AC; OSRAM, Munich, Germany), and a Peltier-cooled scientific charge-coupled device digital camera system (AxioCam HR) driven by the AxioVision 3.0.6.36 software package (Carl Zeiss Vision GmbH). Table 1 summarizes the filter sets used. Unlabeled cells were used to examine autofluorescence. For comparative studies, all photomicrographs in a series of experiments were taken and processed under identical conditions for a given fluorophore.

Colocalization experiments Colocalization experiments were conducted using a Nikon inverted microscope (Eclipse 300TE; Nikon, Vienna, Austria) equipped with a CFI Plan Fluor 40x oil-immersion objective (numerical aperture 1.3; Nikon), an epifluorescence system (150W XBO; Optiquip, Highland Mills, NY), a computer-controlled z-stage (Ludl Electronic Products, Hawthorne, NY), and a liquid-cooled charge-coupled device camera (–30°C; Quantix KAF1400G2; Roper Scientific, Acton, MA). Image resolution was 0.171 µm/pixel. Excitation wavelengths were selected using a computer-controlled filter wheel (Ludl Electronic Products). Deconvolution imaging was performed using ImagePro 3.0 software (Media Cybernetics, Silver Spring, MD). For image analysis and two-dimensional (2-D) deconvolution, Metamorph 6.0 (Universal Imaging, Visitron Systems, Puchheim, Germany) was used.

To identify the compartment that accumulates BODIPY-PGPE, we stained cells for 30 min with three specific organelle markers: a lysosomal marker (LysoTracker Blue DND-22; 160 nM), an ER-Tracker (ER-Tracker Blue-White DPX; 200 nM), or a mitochondrion-selective probe (MitoTracker Red CMXRos; 200 nM), in serum-free culture medium followed by two washing steps with PBS and treatment with 10 nM BODIPY-PGPE for 15 min at 37°C. After rinsing the cells twice with PBS, the coverslip was transferred into an observation chamber and viewed with the fluorescence microscope.

Colocalization studies with BODIPY-POVPE were performed with FM 1-43 membrane probe, which is believed to insert into the outer leaflet of the cell membrane, where it becomes intensely fluorescent. For this purpose, cells were first incubated with 10 nM BODIPY-POVPE for 15 min at 37°C. After a double washing step with PBS, the fluorescence pattern of the labeled phospholipid was imaged in an observation chamber with the fluorescence microscope. Removal of the incubation medium was followed by treatment of the same cells with 150 nM FM 1-43 in prewarmed supplemented DMEM without FCS for 30 min. Finally, cells were thoroughly rinsed with PBS and additional images were acquired from the same subpopulation described above.

For visualization, cells were illuminated at 485 nm (485DF15; Omega Optical), 380 nm (380HT15; Omega Optical), and 575 nm (575DF25; Omega Optical). Emission was monitored at 535 nm (535WB40; Omega Optical) for the BODIPY phospholipids, at 633 nm (528-633DBEM; dichroic XF53; Omega Optical) for the MitoTracker and FM 1-43, and at 510 nm (510WB40; Omega Optical) for the LysoTracker and ER-Tracker.

After staining with the ER-Tracker and LysoTracker, the viability of cells was assessed using 5 µM YO-PRO-1 iodide (Molecular Probes) in serum-free culture medium for 10 min at room temperature. Cytotoxic effects were not observed with these organelle trackers or under the influence of 0.1 µM BODIPY-PGPE at 37°C for 1 h. YO-PRO-1 fluorescence was viewed with identical microscope settings as for BODIPY.

Labeling of cell proteins with BODIPY-POVPE
For each experiment, a fully confluent monolayer of A7r5 cells in one Petri dish (100 mm) was incubated in the dark for 30 min with 8 ml of a 2.5 µM BODIPY-POVPE dispersion in PBS supplemented with 100 mg/l CaCl₂ and 100 mg/l MgCl₂·6H₂O. After treatment, cells were washed twice with ice-cold PBS and then scraped into 3 ml of PBS supplemented with 100 mg/l CaCl₂, 100 mg/l MgCl₂·6H₂O, 1 mM PMSF, and 50 µM NaCNBH₃. The following steps were performed at 4°C. Cells were pelleted at 300 g for 3 min followed by lysis in 150 µl of neutral cell lysis buffer (20 mM HEPES, 2 mM EDTA, 1% Triton X-100, pH 7.4, 50 µM NaCNBH₃, 5 mM DTT, 1 mM PMSF for 1 h on ice; stock solutions of soybean trypsin inhibitor and leupeptin were added directly before use to adjust a final concentration of 10 µg/ml). The suspension was shaken vigorously every 15 min. To remove nuclei and cytoskeleton, the lysate was centrifuged at 1,000 g and 4°C for 5 min. A 1 mM solution of NaCNBH₃ in water was added to the supernatant to reach a final concentration of 100 µM. After reduction of formed Schiff bases at 37°C for 2 h, samples were stored at –20°C. Measurements of cell protein content were performed using a microtiter plate assay based on the method of Bradford (30).

Two-dimensional gel electrophoresis of labeled proteins
For each gel (20 x 20 cm), 200 µg of protein was precipitated according to Wessel and Fluegge (31). The protein pellet was resolubilized in 340 µl of rehydration buffer [7 M urea, 2 M thiourea, 4% CHAPS, 0.002% (w/v) bromophenol blue, and 2% Pharmalyte^TM 3–10] for 30 min at 37°C. Sample solutions were applied to 18 cm immobilized pH gradient strips (Immobiline DryStrip pH 3–10 NL; Amersham Biosciences) (32) by rehydration loading (33, 34). Isoelectric focusing and SDS-PAGE were performed as described previously (35, 36) using 10% polyacrylamide gels. After isoelectric focusing, Immobiline Dry Strip pH 3–10 NL strips were stored at –70°C. Gels were run in a PROTEAN® II XI Multi-cell 2-D apparatus (Bio-Rad) at 10°C, maintaining a limiting current of 50 mA/gel. After electrophoresis, gels were fixed in fixing solution (10% EtOH, 7% AcOH) for at least 2 h. Proteins were detected with the Molecular Imager® FX Pro Plus scanner (at a resolution of 100 µm) driven by PDQuest version 7.1.0 software (both from Bio-Rad). BODIPY fluorescence was determined using an external laser (Bio-Rad) at 488 nm for excitation and a 530/30 nm band-pass filter for the detection of emission. Total protein patterns were visualized by SYPRO Ruby Protein Gel Stain (Molecular Probes) according to the manufacturer's protocol. Scanning parameters for the SYPRO Ruby detection were the same as described previously, except that a 605 DF50 filter was used as the emission filter.

Stability of BODIPY-labeled phospholipids in cells
Confluent monolayer cultures of A7r5 cells in 100 mm Petri dishes were treated with the BODIPY analogs of PGPC (0.1 µM), POVPC (0.5 µM), or POPC (0.1 µM) in 10 ml of serum-free culture medium at 37°C for 5, 15, 30, 45, and 60 min. After incubation, cells were washed once with 10 ml of ice-cold PBS, scraped into 6 ml of ice-cold PBS with a rubber policeman (Corning), and then transferred into a pyrex tube. The culture dish was rinsed with an additional 3 ml of ice-cold PBS. The combined cell suspensions were centrifuged at 300 g and 4°C for 3 min. After discarding the supernatant, cellular lipids were extracted by a simplified procedure of Folch, Lees, and Sloane Stanley (37). The cell pellet was resuspended in 3 ml of CHCl₃/MeOH (2:1, v/v) and was shaken vigorously for 1 h at 4°C. After washing with 700 µl of a MgCl₂ solution (0.036% in water) and centrifuging at 80 g for 2 min at room temperature, the lower chloroform phase was collected and evaporated to dryness. In the case of BODIPY-POVPE, extracted lipids were reduced with 10 µM NaCNBH₃ in a biphasic system (a mixture of 1 ml of diethylether and 1 ml of buffer containing 0.75 M NaCl and 1 mM HEPES) (38), leading to the conversion of formed Schiff base adducts to stable amine structures. The reduced lipids were extracted from the aqueous phase with 5 ml of CHCl₃/MeOH (2:1, v/v). Finally, the isolated lipid mixtures were subjected to TLC (silica gel 60; Merck) using CHCl₃/MeOH/water (65:25:4, v/v/v) as a developing solvent. BODIPY-PGPE, -POVPE, and -POPE served as references (R_f = 0.27, 0.32, and 0.59, respectively). Fluorescence intensities of single spots were determined with a charge-coupled device camera (Herolab, Vienna, Austria) at an excitation wavelength of 365 nm using EasyWin® software for data acquisition.

Two-dimensional TLC separation of labeled cellular lipid extracts
A7r5 cells at high confluence in 100 mm Petri dishes were treated with 1 µM BODIPY-POVPE in 10 ml of serum-free culture medium for 30 min at 37°C. Lipid extraction was performed as described above. Reduction of the imines to secondary amines was carried out according to Friedman et al. (39) in the presence of 10 µM NaCNBH₃. Isolated lipids corresponding to 3–6 µg of phosphorus were applied onto a TLC plate (silica gel 60; Merck) (10 x 10 cm) and subjected to 2-D TLC separation using CHCl₃/MeOH/25% NH₃ (65:25:5, v/v/v) and CHCl₃/acetone/MeOH/glacial AcOH/water (50:20:10:10:5, v/v/v/v/v) as solvents in the first and second directions, respectively. Separated lipids were visualized by iodine vapor after the detection of fluorescent spots with a charge-coupled device camera as mentioned above.

Determination of acid sphingomyelinase activity
Rat aortic smooth muscle cells (A7r5) were grown on six-well plates until 80% confluency was reached. Growth was arrested with DMEM + 0.1% FCS for 24 h, and cells were subsequently incubated with BODIPY-PGPE, BODIPY-POVPE, POVPC, or PGPC (10 µM each) for 30 min. After treatment of the smooth muscle cells with the respective compounds, cells were washed with cold PBS, scraped, harvested, and lysed by incubation with acid lysis buffer for the determination of acid sphingomyelinase activity using a fluorescent sphingomyelin substrate as described previously (26). Aliquots of cell lysates containing 1 µg of protein were analyzed for enzyme activity.

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Synthesis and characterization of fluorescent PGPC and POVPC analogs
Chemical synthesis of seven fluorescently labeled phospholipids carrying a covalently bound BODIPY, Alexa647, or pyrene fluorophore was accomplished to study their interactions with VSMCs. The BODIPY- and Alexa647-labeled PGPC derivatives were prepared from PGPC by PLD-catalyzed head group exchange followed by N-acylation using BODIPY-SE (Fig. 1A) and Alexa647-SE, respectively. Esterification of 1-palmitoyl-sn-glycero-3-phosphocholine with 5,5-dimethoxypentanoic acid, subsequent exchange of choline against ethanolamine, covalent attachment of a BODIPY label to the head group, and final deprotection of the aldehyde group yielded BODIPY-POVPE (Fig. 1B). Synthesis of BODIPY-POPE was accomplished by the acylation of POPE with BODIPY-SE. Pyrene analogs of PGPC, POVPC, and POPC were obtained starting from 1,2-bis(10-pyrenedecanoyl)-sn-glycero-3-phosphocholine, which was converted to the corresponding 2-lysophospholipid by PLA₂. Acylation of this compound with glutaric anhydride or oleic acid completed the synthesis of PyrGPC (Fig. 1C) or PyrOPC, respectively. Condensation of 1-(10-pyrenedecanoyl)-sn-glycero-3-phosphocholine with 5,5-dimethoxypentanoic acid delivered a stable precursor of PyrOVPC, which was deprotected by carefully controlled hydrolysis (Fig. 1C). For more details, see Experimental Procedures.

The new BODIPY and pyrene derivatives were characterized by MALDI-TOF, as shown in Fig. 2 and Table 2 . In all cases, the expected molecular and pseudomolecular ions, respectively, were found. Simple replacement of cations gives the interpretation for the additional signals of (pseudo)molecular ions that are listed in Table 2.

View larger version (17K): [in this window] [in a new window]   Fig. 2. Matrix-assisted laser desorption/ionization time-of-flight (MALDI-TOF) mass spectra of the synthesized BODIPY and pyrene derivatives. The graphs show expanded views of the important parts of the MALDI-TOF mass spectra. Compare Table 2 and see text for discussion. Matrix cluster ions are marked with asterisks. A: BODIPY-POVPE. B: BODIPY-PGPE. C: BODIPY-1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine (POPE). D: PyrOVPC. E: PyrGPC. F: 1-(10-Pyrenedecanoyl)-2-oleoyl-sn-glycero-3-phosphocholine (PyrOPC).

Fluorescence spectroscopic characterization of the labeled phospholipids yielded an excitation maximum of 505 nm and maximal emission at 510 nm for all BODIPY-labeled samples independent of the molecular species. As further summarized in Table 1, phospholipids carrying a pyrene fluorophore uniformly showed excitation and emission maxima at 342 and 376 nm, respectively. Thus, it can be concluded that coupling of either marker to the desired compounds did not significantly affect their spectral properties.

Similar structural tendencies become apparent when tentative space-filling models of POVPC and its pyrene and BODIPY analogs are compared (Fig. 3 ). The compounds shown have in common a large head group including the polar sn-2 substituent and only a single hydrophobic chain.

View larger version (28K): [in this window] [in a new window]   Fig. 3. POVPC and its fluorescent analogs share essential structural features. Tentative space-filling models of POVPC (A), PyrOVPC (B), and BODIPY-POVPE (C) are shown (SYBYL software). Note the high similarity of the natural lipid and its derivatives, especially the pyrene analog, all of them sharing a single long hydrophobic chain and a large polar head group.

In this study, we used BODIPY- and pyrene-labeled lipid analogs for the investigation of lipid import into the cells to detect potential effects of the fluorophore. Some differences were found between the fluorescence images of the PGPC analogs (15 min incubation) and the POVPC analogs at all incubation times. In this context, it has to be emphasized that pyrene fluorescence is much weaker and, in addition, is blue-shifted relative to the emission of BODIPY. As a consequence, the pyrene images are distorted by background fluorescence to a much larger extent. For the same reasons, the tiny contours of plasma membrane fluorescence are seen only with BODIPY but not with pyrene.

Significant intracellular accumulation of blue fluorescence in the perinuclear region was observed after incubation with PyrGPC (Fig. 4A ) relative to untreated control cells (Fig. 4D), even after a period as short as 5 min. This effect persisted for the whole duration of treatment, leading to a steady increment of fluorescence intensity in this intracellular location. A slight staining of the plasma membrane became apparent after 15–30 min but was not further intensified during longer incubation times. The staining pattern of PyrGPC was very similar to that seen when cells were treated with BODIPY-PGPE. At early time points, only regions in close proximity to the cell nucleus fluoresced brightly, whereas later, weak surface labeling occurred and increased in intensity. Under the same experimental conditions, Alexa647-PGPE partitioned in an analogous manner after 5 and 30 min.

View larger version (78K): [in this window] [in a new window]   Fig. 4. Time-dependent uptake of fluorescent PGPC, POVPC, and POPC analogs into vascular smooth muscle cells (VSMCs). A–C: A7r5 cells were incubated with 0.1 µM BODIPY-PGPE, 2 µM PyrGPC, or 0.1 µM Alexa647-PGPE (A), 0.5 µM BODIPY-POVPE or 2 µM PyrOVPC (B), and 0.1 µM BODIPY-POPE or 2 µM PyrOPC (C) in serum-free culture medium for 5, 15, 30, or 45 min at 37°C, washed twice with PBS, and observed by fluorescence microscopy. D: Autofluorescence micrographs of VSMCs treated with culture medium without fetal calf serum (FCS) for 5–60 min. No differences could be observed for varying incubation periods. Using the optical settings for Alexa647, absolutely no background was detected. Photomicrographs shown are representative of at least five cells for each time point in three or more independent experiments, with the exception of the Alexa647 images (single experiment). For a given microscope channel, all photomicrographs were exposed and printed identically. Bars = 20 µm. Fluorescent derivatives of both oxidized phospholipids are rapidly internalized by VSMCs, in contrast to their POPC equivalents. Note the remarkable differences in their intracellular distribution. PGPC analogs are initially transferred to perinuclear regions, whereas in POVPC derivatives, fluorescence becomes highly enriched in the plasma membrane during short incubation times. There is a high correlation between photomicrographs obtained from each phospholipid species independent of the type and the binding site of the respective label.

View larger version (77K): [in this window] [in a new window]   Fig. 5. Colocalization of BODIPY-PGPE and BODIPY-POVPE with organelle-specific markers in VSMCs. Subcellular distribution of BODIPY-PGPE in VSMCs with respect to various organelle-specific markers. A–C: A7r5 cells were pretreated with LysoTracker Blue DND-22 (160 nM) (A), ER-Tracker Blue-White DPX (200 nM) (B), or MitoTracker Red CMXRos (200 nM) (C) for 30 min, followed by incubation with 10 nM BODIPY-PGPE in supplemented DMEM without serum for 15 min. Fluorescence images of both the BODIPY phospholipid and the organelle-specific probes were recorded from the same field of cells and then used to produce overlays of the double staining. Note the extensive colocalization of BODIPY and LysoTracker fluorescence. Although partial delivery of the labeled lipid to the endoplasmic reticulum (ER) is also evident, mitochondria can be excluded as an intracellular destination of exogenously supplied BODIPY-PGPE under the experimental conditions used. D: Treatment of A7r5 cells with BODIPY-POVPE in serum-free culture medium for 15 min and image acquisition preceded incubation with FM 1-43 membrane probe (150 nM) in fresh medium without FCS for 30 min. The field previously imaged for BODIPY fluorescence was rephotographed for FM 1-43 fluorescence. The extensive overlap of the dual-labeled structures suggests a predominant accumulation of BODIPY-POVPE in the plasma membrane of VSMCs. Photomicrographs shown are representative of at least three independent experiments. In control experiments, hardly any crossover between green (BODIPY-PGPE), blue (LysoTracker, ER-Tracker), and red (MitoTracker, FM 1-43) fluorescence channels was detected at these concentrations. For a given microscope channel, all photomicrographs were exposed and printed identically. Bars = 20 µm.

PyrOVPC basically showed the same fluorescence pattern as its BODIPY analog. However, the plasma membrane staining after long incubation times was less pronounced. This is very likely attributable to the fact that pyrene at sufficiently high concentrations in the bilayer may form excited-state dimers (excimers) upon excitation emitting at 470 nm, although with a lower intensity compared with the blue monomer (44).

Fluorescent POPC analogs (pyrene- or BODIPY-labeled) as nonoxidized reference compounds with two long hydrophobic chains were not efficiently taken up by the cells in the same time domain and, in addition, showed a different intracellular distribution pattern (Fig. 4C). In summary, it is quite obvious that short-chain oxidized phospholipids partition into cells very fast and are targeted to different subcellular compartments compared with the more hydrophobic double-chain phospholipids.

Uptake of pyrene-labeled POVPC analogs depends on LDL internalization
In addition to pure dispersions of oxidized phospholipids, we used LDL as a delivery system for the import of these compounds into cells. When A7r5 cultures were incubated with LDL (50 µg protein/ml) containing PyrGPC (3 nmol/µg protein), a significant amount of the labeled lipid was already transferred to cells after 15 min (Fig. 6A ). The pattern of intracellular fluorescence was similar to that observed previously using pure PyrGPC dispersions (Fig. 4A). Longer incubations (45 min) led to an additional accumulation of blue fluorescence in internal structures around the nucleus, with only a tiny fraction being located in the plasma membrane. In contrast, different images were obtained when cells were treated with LDL (50 µg protein/ml) preloaded with PyrOVPC (5 nmol/µg protein) compared with pure lipid aggregates as donor systems. Surface labeling still occurred, but the amount of internalized lipid was much higher and the distribution pattern was different (Fig. 6B). Obviously, the LDL particles change the (unknown) mechanism of translocation and distribution of the fluorescent POVPC analog in VSMCs.

View larger version (30K): [in this window] [in a new window]   Fig. 6. Transfer of PyrGPC and PyrOVPC from LDL into VSMCs. A7r5 cells were incubated with LDL/PyrGPC (50 µg protein/ml; 3 nmol PyrGPC/µg protein) (A), LDL/PyrOVPC (50 µg protein/ml; 5 nmol PyrOVPC/µg protein) (B), BODIPY-LDL/PyrGPC (50 µg protein/ml; 3 nmol PyrGPC/µg protein) (C), and BODIPY-LDL/PyrOVPC (50 µg protein/ml; 5 nmol PyrOVPC/µg protein) (D) in serum-free culture medium for 15 or 45 min at 37°C, washed twice with PBS, and observed with the fluorescence microscope. Images were acquired at blue (pyrene) and green (BODIPY) wavelengths. Photomicrographs shown are representative of at least three cells for each time point in three independent experiments. In control experiments, hardly any crossover between green (BODIPY) and blue (pyrene) wavelengths was detected at these concentrations. For a given microscope channel, all photomicrographs were exposed and printed identically. Bars = 20 µm. Similar uptake rates and fluorescence patterns were observed for PyrGPC and PyrOVPC when delivered from single (A, B) or double (C, D) labeled LDL, indicating no interference of the attached BODIPY fluorophore with the phospholipid exchange process. Note that the intracellular fluorescence pattern obtained after treatment with LDL, which was prelabeled with pyrene analogs of either oxidized phospholipid, highly resembles the distribution observed when cells were incubated with aqueous dispersions of fluorescent PGPC derivatives (see Fig. 4A, B). The absence of green intracellular labeling (C, D) indicates no uptake of LDL by VSMCs under these experimental conditions. apoB, apolipoprotein B-100.

Stability of BODIPY-labeled PGPC and POVPC analogs in cells
Lipid extraction of cells after incubation with BODIPY-PGPE followed by TLC analysis showed limited stability of this compound in VSMCs. A new fluorescent lipid was formed corresponding to the respective lysophospholipid on TLC (Fig. 7 ). Experiments to determine the stability of the pyrene-labeled PGPE under reasonable conditions relevant to the fluorescence microscopy studies (0.1 mM lipid concentration) failed. Pyrene fluorescence is lower than BODIPY fluorescence by at least 1 order of magnitude and, in addition, blue-shifted. Thus, it is not detectable in the background fluorescence of the lipid extract. According to all we know from lipids labeled with pyrene in the acyl chain (minimum perturbing effect) (45), these compounds are better substrates for enzymes. Therefore, pyrene-PGPE is likely to be hydrolyzed faster inside the cells compared with the BODIPY derivative.

View larger version (10K): [in this window] [in a new window]   Fig. 7. Degradation of BODIPY-PGPE in VSMCs. A7r5 cells were incubated for the indicated times with 0.1 µM BODIPY-PGPE in serum-free medium, washed, and collected. After lipid extraction and TLC separation, BODIPY fluorescence was quantified directly on the TLC plates. BODIPY-PGPE was continuously hydrolyzed to BODIPY-phosphoethanolamine (lysoPE), which constitutes the only newly formed fluorescent spot. Data are representative of two independent experiments producing similar results.

View larger version (25K): [in this window] [in a new window]   Fig. 8. Detection of modified lipids in VSMCs by covalent labeling with BODIPY-POVPE. Total lipid extracts from A7r5 cells were preincubated with 1 µM BODIPY-POVPE in serum-free culture medium (see Experimental Procedures). After reduction with 10 µm NaCNBH3, they were separated by two-dimensional (2-D) TLC using CHCl3/MeOH/25% NH3 (65:25:5, v/v/v) and CHCl3/acetone/MeOH/glacial AcOH/water (50:20:10:10:5, v/v/v/v/v) in the first and second direction, respectively. The starting point was the lower left corner. A: Lipid products with the fluorescent phospholipid were detected with a charge-coupled device camera as described in Experimental Procedures. Arrows indicate fluorescent spots most likely corresponding to modified cellular lipids, presumably phosphatidylethanolamine (PE) and phosphatidylserine (PS). The arrowhead denotes intact BODIPY-POVPE. The remaining fluorescent spots probably represent self-condensation products of the same compound. B: Iodine vapor was used as a general lipid stain. However, unsaturated lipids are more intensely stained. This is probably the reason why most of the fluorescent spots apart from the presumed phosphatidylethanolamine product could hardly be visualized by this method, as they consist mainly of BODIPY-POVPE containing only a single saturated fatty acyl chain. PC, phosphatidylcholine; PE, phosphatidylethanolamine; PI, phosphatidylinositol; PS, phosphatidylserine.

View larger version (23K): [in this window] [in a new window]   Fig. 9. Detection of primary protein targets in VSMCs by covalent labeling with BODIPY-POVPE. Total protein extracts from A7r5 cells preincubated with 2.5 µM BODIPY-POVPE in PBS (see Experimental Procedures) were separated by 2-D gel electrophoresis after reduction with 70 µM NaCNBH3. A: Protein adducts with the fluorescent phospholipid were detected with a laser scanner as described in Experimental Procedures. B: SYPRO Ruby Protein Gel Stain was used to visualize total protein. Note that not all but still numerous proteins, including low-abundance proteins, became visible under the experimental conditions used, indicating covalent modification by BODIPY-POVPE.

View larger version (13K): [in this window] [in a new window]   Fig. 10. Effects of BODIPY-labeled and unlabeled oxidized phospholipids on acid sphingomyelinase activity in VSMCs. Rat aortic smooth muscle cells (A7r5) were incubated with BODIPY-PGPE, BODIPY-POVPE, POVPC, or PGPC (10 µM each) for 30 min. Cells were isolated and lysed in lysis buffer for the determination of acid sphingomyelinase activity (see Experimental Procedures). Enzyme activities were measured as described previously (26) and expressed as percentage of control (Co). Data are means ± SD (n = 3). * P < 0.05 compared with control.

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

To exclude or better estimate the influence of the lipid-bound fluorophore, we synthesized up to three differently labeled analogs to verify the relevance of our observations. In these approaches, we set a high value on conforming size and polarity of the marker to match the properties of the unlabeled parent compounds as closely as possible. The BODIPY fluorophore was chosen by virtue of its high fluorescence yield and great photostability. Because of its rather polar nature, it was linked to the head group of the phospholipid molecules. The same applies to the even more polar Alexa647 label, which has proven to be particularly suitable for minimizing background fluorescence. In contrast, the spectral properties of pyrene analogs do not allow for high-sensitivity detection, but they have the advantage of carrying a hydrophobic label at the terminus of the long sn-1 acyl chain. If embedded in a membrane, the marker will most likely reside in the very center of the bilayer, where the molecular order and, thus, membrane perturbation should be minimal (45).

Our fluorescence microscopy experiments indicate high agreement between data obtained from the same phospholipid species containing different labels. Thus, it may be concluded that the observed phenomena can be attributed to innate properties of the lipid molecules rather than to the attached fluorophores. This is not surprising, because the fluorescent derivatives and their parent compounds share basic structural features. As tentatively illustrated in Fig. 3, they are characterized by a single hydrophobic chain and an oversized polar head group, both of which in combination confer high surface activity. Taking this into consideration, it can be expected that oxidized phospholipids are rapidly exchanged between membranes and/or liposomes. Previous studies have demonstrated that, when cells or lipid vesicles are incubated with liposomes containing another short-chain phospholipid, 1-acyl-2-(N-4-nitrobenzo-2-oxa-1,3-diazole)-aminocaproyl phosphatidylcholine, the fluorescent lipid spontaneously transfers between membranes (47). The mechanism for this transfer has been demonstrated to be diffusion of soluble fluorescent lipid monomers (48). It is very likely that the exchange of the even more polar oxidized phospholipids under investigation between the donor particles and cells is driven by the same process. Moreover, the oxidized phospholipids are probably monomeric, as lysolecithin, whose polarity resembles that of oxidized phospholipids, has a critical micelle concentration of 450 ± 60 µM in aqueous solution at 25°C (49), far higher than the concentration of the oxidized lipids under the incubation conditions used (0.1–2 µM).

Indeed, fluorescence microscopy revealed very fast uptake of synthetic PGPC analogs by VSMCs, resulting in a lysosomal distribution pattern (Fig. 5A). It was not until 30 min that a slight but distinctive staining of the plasma membrane became visible as well. The pathway by which intracellular membranes are labeled by PGPC derivatives is unknown. One possible explanation is that intact PGPC derivatives are targeted directly to the lysosomes, reflecting either vesicular movement or monomeric diffusion. Alternatively, staining of ER membranes may be the result of lipid degradation. The latter possibility could be ruled out by the finding that the half-time of BODIPY-PGPE in VSMCs is >1 h. Figure 7 shows the partial hydrolysis of BODIPY-PGPE to the respective lysolipid as the only fluorescent metabolite newly formed. This observation gives rise to the notion that the apoptotic signaling, which has been shown to be triggered by PGPC within 10 min (11), can be ascribed without much doubt to intact molecules of the oxidized phospholipid. Nevertheless, it cannot be excluded that the repartition of fluorescence during prolonged incubations originates from the anterograde translocation of cumulative hydrolytic degradation products.

The converse situation, in which prominent staining of the plasma membrane clearly precedes an intracellular increment of fluorescence intensity, was observed when cells were incubated with the aldehydolipid (POVPC) derivative for short periods of time (Fig. 4B), perhaps as a consequence of covalent reaction with amino and other groups in lipids and proteins. Previous reports already demonstrated that other aldehydes, particularly 4-hydroxy-2-nonenal (HNE), are able to react with amino groups of lysines and histidines and with sulfhydryl groups of proteins and peptides, leading to the formation of HNE adducts with cellular proteins during oxidative stress (50, 51). Of great relevance to this work is the fact that HNE can activate the epidermal growth factor receptor and its downstream signaling cascades merely through derivatization of reactive amino groups (52, 53). On the other hand, aldehydes generated by the myeloperoxidase system of phagocytes were shown to covalently modify the amino groups of ethanolamine and serine glycerophospholipids to form biologically active Schiff base adducts (46). Zieseniss et al. (38) found that Schiff base adducts between phosphatidylethanolamine and aldehydes in oxidized LDL provoke the stimulation of platelet prothrombinase activity. In the beginning, there was little consideration of the possibility that some of the masking of amino groups during LDL oxidation might be attributable to conjugation with phospholipid aldehydes, as exemplified by POVPC. Several studies now exist that demonstrate covalent adduct formation between oxidized phospholipids and apoB (13, 38, 54). In this work, we found evidence that BODIPY-POVPE reacts with many cellular proteins. Multiple fluorescent protein spots could be detected upon separation of prelabeled cell protein extracts by 2-D gel electrophoresis (Fig. 9). In addition, 2-D TLC revealed that at least one fraction of cell lipid was also covalently modified by BODIPY-POVPE (Fig. 8). In summary, it seems possible that POVPC is inserted rapidly into the plasma membrane of VSMCs during its first contact with the cell surface, resides there in covalent association with amine-containing lipids and proteins, and is subsequently, at least in part, transferred in the form of Schiff bases to perinuclear regions.

Another interesting point pertaining to the current study is the finding that the way that oxidized phospholipids are presented to VSMCs has some influence on their uptake rates and intracellular distribution. LDL enriched with the pyrene analogs of PGPC yielded a fluorescence pattern similar to that observed when cells were treated with aqueous dispersions of the respective lipids. In contrast, the import of LDL-associated pyrene-POVPC was more efficient compared with that of the free lipid. Experiments with dual-labeled LDL (BODIPY at apoB, green fluorescence; pyrene-labeled lipids, blue fluorescence) produced the same results. All experiments were conducted with nonfasted cells, which means that LDL receptor expression at the cell surface was repressed (55). Thus, LDL internalization is only partially involved in the observed effects, and lipid import takes place via lipid transfer to a large extent. Obviously, the polar truncated phospholipids are easily exchangeable between different lipoprotein particles and/or cellular membranes. In this way, they can broaden their cross-section of action beyond a single cell or particle, thus exerting biological activity distant from their sites of origin. This idea is in agreement with the results of Podrez et al. (56), who suggested that oxidized phospholipids not adducted to apoB within oxidized LDL can transfer to native LDL and induce CD36 recognition by macrophages. By this means, they can be considered important mediators of cytotoxic effects (11) elicited by oxidative stress in other parts of biological systems as well. For instance, the possibility that oxidized phospholipids are transferable between oxidized LDL and intact HDL may lead to multiple functions in the development of atherosclerosis (57).

In contrast to POVPC, the cellular effects attributable to PGPC are entirely physical. Owing to its excellent membrane-perturbing capacity, it is conceivable that PGPC is capable of altering lipid packing, perhaps not only in fluid but also in more rigid microdomains such as rafts or caveolae (58). Many receptor-mediated signaling systems may be localized to and operate through caveolae, including epidermal growth factor receptor, Ras, and trimeric G-proteins, among others (59). Any membrane protein, but particularly ion channels, may be regulated via the cellular control of membrane composition, as indicated by the responsiveness of specific ion channels to changes in the lipid environment (60). Differences in the biophysical properties of membrane microdomains (e.g., fluidity) have been suggested to play a role in the sorting of proteins and lipids into different subpopulations of intracellular transport vesicles (61, 62), possibly resulting in abnormal trafficking. Thus, the physical interaction of PGPC with cellular components or the deregulation of key signaling pathways that probably operate through membrane microdomains could lead to aspects of the differentiated VSMC phenotype.

In general, the physical interaction and covalent modification of cellular components by oxidized phospholipids exhibit only poor specificity, which suggests the involvement of various mediators and the induction of a wide range of biological effects. PGPC and POVPC have been reported to elicit apoptotic signaling in VSMCs (11). Issues that remain to be clarified include the identification of their primary molecular targets. Here, we demonstrate the presence of multiple interaction sites for oxidized phospholipids on the cell surface as well as inside VSMCs. It is intriguing to consider the possibility that signal transduction pathways might be activated simultaneously at different levels, even bypassing early signaling elements.

Walton et al. (19, 63) suggested that the induction of interleukin-8 transcription by oxidized 1-palmitoyl-2-arachidonoyl-sn-glycero-3-phosphocholine is mediated by a 37 kDa glycosylphosphatidylinositol-anchored protein that interacts with Toll-like receptor 4. On the basis of studies in Xenopus laevis oocytes, Leitinger and colleagues (64) concluded that POVPC and PGPC elicit their cellular responses through binding to different receptors. This adds to earlier indications that the effects of oxidized phospholipids are receptor-mediated. Abolishment of their cellular responses by platelet-activating factor receptor antagonists implies a central role for this receptor in activating VSMCs, endothelial cells, monocytes, platelets, and neutrophils (10, 65–67). Subbanagounder et al. (68) have provided evidence that the in vitro effects of POVPC and 1-palmitoyl-2-(5,6-epoxyisoprostane E₂)-sn-glycero-3-phosphocholine, another bioactive lipid present in minimally modified LDL (69), probably are not mediated by the platelet-activating factor receptor. They do not exclude a role for it in the action of these two molecules and speculate that there might exist a separate WEB 2086-sensitive receptor for POVPC and 1-palmitoyl-2-(5,6-epoxyisoprostane E₂)-sn-glycero-3-phosphocholine. We acknowledge the involvement of membrane proteins in particular processes activated by oxidized phospholipids, but in the light of the data presented here, we doubt the exclusive role of specific receptors for these compounds and support the notion that the majority of the observed effects result from many different interactions with components of their microenvironment. This idea is consistent with the results of Kogure et al. (23) suggesting that apoptosis induced by oxidized LDL-derived phospholipids in VSMCs is caused by temporary membrane distortion and not through specific receptors.

Further studies are needed to elucidate the mechanisms underlying the pathophysiological effects of oxidized phospholipids on VSMCs. The identification of their primary molecular targets is the subject of intense investigation. The knowledge of all of the signal and receptor components potentially involved in the functional deregulation of vascular cells by oxidized lipid species will have a profound impact on understanding and modulation of atherogenesis.

	ACKNOWLEDGMENTS

 
The authors thank Mr. Stefan Weißgerber for exceptional help with image processing. The authors are grateful to Dr. Dieter F. Münzer for valuable assistance with the SYBYL software. The authors acknowledge financial support from the Austrian Science Fund (FWF projects P14714-PHA, P13962-CHE, and SFB F30-B05).

Manuscript received February 1, 2006 and in revised form November 29, 2006.

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