Apolipoprotein E Inhibits Platelet-derived Growth Factor-induced Vascular Smooth Muscle Cell Migration and Proliferation by Suppressing Signal Transduction and Preventing Cell Entry to G1 Phase*

Masato Ishigami, Debi K. Swertfeger, Norman A. Granholm, and David Y. HuiDagger

From the Department of Pathology and Laboratory Medicine, University of Cincinnati College of Medicine, Cincinnati, Ohio 45267-0529

    ABSTRACT
Top
Abstract
Introduction
Procedures
Results
Discussion
References

The anti-atherogenic effects of apolipoprotein (apo) E have been attributed to its ability to reduce plasma cholesterol level and to limit foam cell formation. The purpose of this study was to ascertain if apoE also may have cytostatic functions that could attenuate vascular occlusive diseases. Purified apoE inhibited smooth muscle cell migration directed to platelet-derived growth factor (PDGF) or oxidized LDL (oxLDL) (p < 0.0001). The purified apoE also suppressed PDGF- and oxLDL-induced smooth muscle cell proliferation (p < 0.001). These apoE inhibitory effects were not because of suppression of PDGF binding to its receptors on the smooth muscle cells, but was correlated with a significant reduction in agonist-stimulated mitogen-activated protein kinase activity (p < 0.01). ApoE also inhibited PDGF-induced cyclin D1 mRNA expression, suggesting that the apoE effect was mediated by growth arrest at the G0 to G1 phase. Taken together, these results suggest that apoE has cytostatic functions in the vessel wall and may protect against vascular diseases through inhibition of cell signaling events associated with growth factor-induced smooth muscle cell migration and proliferation.

    INTRODUCTION
Top
Abstract
Introduction
Procedures
Results
Discussion
References

Research in the past two decades has clearly established that apolipoprotein (apo)1 E protects against vascular diseases (1). Early studies suggest that the protective effect may be related to the ability of apoE to promote cholesterol efflux from peripheral cells and to mediate the clearance of cholesterol-enriched lipoproteins from circulation (1). Transgenic mice overexpressing rat apoE displayed marked resistance to diet-induced hypercholesterolemia and did not develop atherosclerosis (2, 3). In contrast, mice with targeted disruption of the apoE gene developed spontaneous atherosclerosis even under basal low fat/low cholesterol dietary conditions (4-7). Atherosclerosis in apoE-null mice could be prevented by increasing circulating apoE level through recombinant adenovirus-mediated apoE gene transfer to the liver (8). The decrease in atherosclerosis in this model was accompanied by decreased total cholesterol and VLDL/IDL levels (8).

The relationship between apoE and cholesterol transport suggests that it prevents atherosclerosis by lowering of plasma cholesterol level. However, atherosclerosis was found to be more severe in chow-fed apoE-null mice than in cholesterol-fed apoE(+/+) mice, despite the relative similar plasma cholesterol level in the two groups (7). Additionally, transgenic expression of apoE in the arterial wall inhibited atheroma formation and severity without affecting plasma cholesterol level and lipoprotein profile in cholesterol-fed C57BL/6 mice (9). Although these two studies characterized atherosclerotic lesions based solely on macrophage foam cell deposition, it is noteworthy that increased number of intimal smooth muscle cells was observed in stenotic lesion area of the external carotid artery (10) and fibroproliferative atherosclerotic plaques of chow-fed apoE-null mice (11). Taken together, these results suggest that apoE may have direct impact on vascular occlusive diseases in a manner in addition to, and independent of, its property as a cholesterol-transporting apolipoprotein. Recently, apoE gene polymorphism has been shown to correlate with risk of restenosis after balloon angioplasty (12). Because lipid deposition and foam cell formation is a late event in restenosis, this correlation adds support to the hypothesis that apoE may have cell regulatory functions in the vessel wall.

A prominent vascular abnormality observed in both spontaneous atherosclerosis and restenosis after angioplasty is the migration of vascular smooth muscle cells from the medial layer of the blood vessel to its intima, followed by proliferation of the intimal smooth muscle cells. These processes are regulated by growth factors released from cells within the injured artery or from circulating cells in plasma. Potent mitogens and chemoattractants that participate in vascular disease include PDGF and oxLDL. These reagents direct migration of vascular smooth muscle cells from the media to the intima of the vessel and induce proliferation of the intimal smooth muscle cells (13, 14). The mechanism of cell activation is reported to be mediated via the activation of mitogen-activated protein (MAP) kinase (15). Because apoE has been shown to inhibit mitogen-stimulated lymphocyte proliferation through suppression of early cell signaling events (16), the current study is undertaken to explore the possibility that apoE may protect against vascular diseases by inhibiting vascular smooth muscle cell migration and proliferation.

    EXPERIMENTAL PROCEDURES
Top
Abstract
Introduction
Procedures
Results
Discussion
References

Materials-- Human recombinant PDGF-BB, FBS, and Dulbecco's modified Eagle's medium (DMEM) were purchased from Life Technologies, Inc. (Gaithersburg, MD). [Methyl-3H]thymidine, [(gamma -32P]ATP, [alpha -32P]dCTP, and 125I-PDGF-BB were obtained from NEN Life Science Products. Activated MAP kinase, insulin-regulated phosphorylated heat- and acid-stable protein (PHAS-I), the MAP kinase assay kit, and QuikhybTM hybridization solution were products of Stratagene (La Jolla, CA). The Cell Titer 96TM AQueous cell proliferation assay kit was purchased from Promega, Inc. (Madison, WI). The embryonic rat aortic smooth muscle-derived A7r5 cells were obtained from American Type Culture Collection (Manassas, VA). Human coronary artery smooth muscle cells were purchased from Clonetics (San Diego, CA). Transwell chambers with gelatin-treated polycarbonate membranes were obtained from Corning Coster Corp. (Cambridge, MA).

Cell Culture-- The A7r5 cells were grown in monolayer cultures on plastic tissue culture-treated multiple well cluster plates in DMEM containing 10% FBS, 100 units/ml penicillin, 0.1 mg/ml streptomycin, and 2 mM glutamine. Cells were incubated at 37 °C in 95% air, 5% CO2 atmosphere and were used for experiments between passages 3-12. Human coronary artery smooth muscle cells were grown in smooth muscle cell growth medium provided by the supplier. Cells were used for experiments between passages 5 and 8. In all experiments, the cells were made quiescent by incubating for 48 h in the presence of 0.4% FBS prior to use.

Preparation of Apolipoproteins and Lipoproteins-- Human apoE was isolated from fresh plasma obtained from normal healthy volunteers as detailed by Rall et al. (17). The purity of the apoE preparation was verified based on a single band with Mr = 34,000 in SDS-polyacrylamide gels. Human LDL (d = 1.019-1.063 gm/ml) was isolated by sequential ultracentrifugation and was stored in saline-EDTA solution (18). Oxidative modification was undertaken by dialyzing the LDL against saline without EDTA and then incubating the lipoproteins with 5 µmol/L copper sulfate for 48 h at 37 °C. The oxidative modification was confirmed by 37 °C electrophoretic mobility on agarose gels.

Smooth Muscle Cell Migration Assay-- The migration of smooth muscle cells was examined according to the procedure described by Law et al. (19). Briefly, the cells were suspended in DMEM with 0.4% FBS at a concentration of 2 × 105 cells/ml. The cells were preincubated with 0, 25, or 50 µg/ml apoE for 30 min at 37 °C, and 0.1-ml aliquots of the cell suspension (2 × 104 cells) were added to the top chamber of gelatin-treated Transwell polycarbonate membrane with 8-µm pores in 24-well plates. The lower Transwell compartment contained 0.6 ml of DMEM, 0.4% FBS, with or without PDGF-BB or oxLDL. Incubation was continued for 4 h at 37 °C, after which the adherent cells were washed extensively, fixed with methanol, and then stained with hematoxylin. The number of cells that migrated to the lower surface of each filter was counted in different high power fields (HPFs) at a magnification of 320. 

[Methyl-3H]Thymidine Incorporation into Cellular DNA-- Cells were plated at a cell density of 5 × 103 cells/well in 96-well tissue culture dishes and then incubated for 48 h in DMEM with 0.4% FBS to synchronize cells at the quiescent state. The cells were incubated for 20 h with PDGF-BB or oxLDL in the presence or absence of apoE. One µCi of [methyl-3H]thymidine was added to the culture medium and the incubation was continued for 4 h at 37 °C. Cells were washed twice with phosphate-buffered saline followed by incubation at 4 °C in 25% trichloroacetic acid for 20 min. The plates were washed three times with cold 25% trichloroacetic acid followed by the addition of 0.25 N NaOH. Radioactivity in the cell lysate was quantitated by liquid scintillation counting.

Cell Proliferation Assay-- Smooth muscle cell proliferation was also assessed based on the ability of the cells to convert MTS into formazan, using the AQueous Cell Proliferation Assay Kit. In these experiments, 5 × 103 quiescent A7r5 or human coronary artery smooth muscle cells in 96-well tissue culture dishes were incubated for 24 h with or without PDGF-BB or oxLDL and in the presence or absence of apoE prior to the addition of 20 µl of MTS mixed with the electron-coupling reagent phenazine methosulfate. Incubation was continued for 1 h at 37 °C. Formazan formation was measured based on increased absorbance at 490 nm.

Binding of PDGF to Rat Aortic Smooth Muscle Cells-- Quiescent A7r5 cells plated in 24-well tissue culture dishes were pre-incubated for 3 h at 4 °C with Hanks' balanced salt solution containing 0.2% bovine serum albumin, with or without addition of 25 or 50 µg/ml apoE or 200 ng/ml PDGF-BB. At the end of this incubation period, 120 pmol/liter of 125I-PDGF-BB was added to each well, and incubation was continued for 3 h at 4 °C. The cells were washed three times with cold Hanks' solution and then solubilized with 1 ml of solution containing 1% Triton X-100 and 0.1% bovine serum albumin. The radioactivity associated with the cell lysate was quantitated in a gamma counter. Specific binding was determined as the total amount of 125I-PDGF-BB bound to the cells minus the nonspecific binding observed in the presence of excess unlabeled PDGF-BB.

Determination of Cyclin D1 mRNA Level in Rat Aortic Smooth Muscle Cells-- A cDNA probe for rat cyclin D1 was prepared by reverse transcription-polymerase chain reaction amplification of mRNA isolated from proliferating cells. The forward oligonucleotide primer (5'-CTGACTGCCGAGAAGTTGTGCATC-3') and the reverse primer (5'-CTGGCTCCTTTCTCTTCGCGATG-3') were designed based on the published sequence of rat cyclin D1 cDNA (20). The 591-base pair amplification product was used for hybridization with 10 µg of cellular RNA extracted using the guanidine thiocyanate-phenol-chloroform method (21). Hybridization was carried out for 3 h at 65 °C with [alpha -32P]dCTP-labeled cyclin D1 and glyceraldehyde-3-phosphate dehydrogenase cDNA probes in QuikhybTM solutions containing denatured salmon sperm DNA. The membrane was washed three times with 1× SSC buffer (1.5 mM sodium citrate, 15 mM NaCl, and 0.1% SDS) followed by another wash with 0.1 × SSC. Hybridization signal was analyzed using the Scion Image PC program. The relative level of cyclin D1 mRNA was determined based on the amount of glyceraldehyde-3-phosphate dehydrogenase mRNA in each sample.

Determination of MAP Kinase Activity-- The smooth muscle cells were plated at a density of 1.5 × 105 cells in 6-well plates and then incubated for 5 min at 37 °C in the presence or absence of PDGF-BB, oxLDL, and apoE. Cell lysate was prepared by adding 200 µl of buffer containing 25 mM HEPES, pH 7.5, 0.2 mM phenylmethylsulfonyl fluoride, 0.05% 2-mercaptoethanol, and 1% Triton X-100. The cell lysate was mixed with 40 µl of kinase buffer containing 25 mM HEPES, pH 7.5, 10 mM magnesium acetate, 50 mM ATP, and 2 µl of [(alpha -32P]ATP (1.0 mCi/ml). The reaction was initiated by adding 10 µl of PHAS-I, and incubation was continued for 10 min at 30 °C. The phosphorylation reaction was terminated by adding 5 µl of 0.25 M phosphoric acid. Ten µl of each reaction mixture was spotted onto chromatography paper, washed with 75 mM phosphoric acid, rinsed in 95% ethanol, and then subjected to liquid scintillation counting.

Statistical Analysis-- The apoE effect on cell proliferation was assessed by two-factorial ANOVA. All other data were analyzed for statistical significance using one-way ANOVA. A p value of less than 0.05 was considered to be statistically significant.

    RESULTS
Top
Abstract
Introduction
Procedures
Results
Discussion
References

ApoE Inhibits PDGF-directed Smooth Muscle Cell Migration-- Using the cell migration assay described under "Experimental Procedures," 6.3 ± 2.6 quiescent A7r5 cells were found to be present in each HPF, indicating that 4.2% of the total amount of plated cells migrated to the lower chamber under basal conditions. The addition of PDGF enhanced A7r5 cell migration 4-fold, with 25.8 ± 6.1 cells observed in each HPF, representing 17.1% of the total amount of plated cells migrated to the lower chamber (Fig. 1A). Human apoE, at 25 µg/ml, inhibited the PDGF-induced A7r5 cell migration to the same level as that observed when the cells were incubated without PDGF (Fig. 1A). Significantly, at 50 µg/ml of apoE, the number of rat aortic smooth muscle cells migrated to the lower chamber was lower than that observed with unstimulated cells (Fig. 1A). These concentrations of apoE have no effect on cell viability as determined by dye exclusion assay. Similar results were observed when experiments were performed with human coronary artery smooth muscle cells instead of the rat A7r5 smooth muscle cells (Fig. 1B). Thus, these experiments demonstrated a role of apoE in inhibiting smooth muscle cell migration with or without growth factor stimulation.


View larger version (17K):
[in this window]
[in a new window]
 
Fig. 1.   Effect of apoE on PDGF-directed migration of smooth muscle cells. Quiescent rat A7r5 smooth muscle cells (panel A) or human coronary artery smooth muscle cells (panel B) were incubated for 30 min at 37 °C with or without apoE and then added to Transwell membranes in 24-well plates at a density of 2 × 104 cells/well. The lower Transwell chamber contained 0.6 ml of DMEM, 0.4% FBS, with or without 10 ng/ml of PDGF-BB. Incubation was continued for 4 h at 37 °C. The number of cells that migrated to the lower surface was determined microscopically by counting in different high power fields (HPF) at a magnification of 320. Data are expressed as mean ± S.D. (n = 6). *, values differed from no apoE; PDGF-stimulated control at p < 0.0001.

ApoE Inhibits PDGF-Stimulated Smooth Muscle Cell Proliferation-- Incubation of quiescent rat A7r5 aortic smooth muscle cells with PDGF resulted in a 2- to 2.5-fold induction of [3H]thymidine incorporation into their DNA in comparison with cells incubated without the growth factor (Fig. 2). The addition of 25 and 50 µg/ml of human apoE to the culture media significantly reduced the PDGF-induced [3H]thymidine incorporation into DNA (p < 0.001, Fig. 2). To confirm that apoE inhibition of PDGF-induced [3H]thymidine incorporation into DNA is a reflection of its ability to inhibit cell proliferation, a direct cell proliferation assay was performed. The results showed that PDGF stimulated the proliferation of both rat embryonic smooth muscle cells and human coronary artery smooth muscle cells in a time-dependent manner (Fig. 3). The addition of apoE effectively inhibited the PDGF-stimulated smooth muscle cell proliferation (p < 0.001, Fig. 3).


View larger version (20K):
[in this window]
[in a new window]
 
Fig. 2.   Effect of apoE on PDGF-induced proliferation of aortic smooth muscle cells. Serum-starved rat A7r5 smooth muscle cells were incubated in 96-well plates (5 × 103 cells/well) for 20 h at 37 °C in DMEM with or without 10 ng/ml of PDGF and apoE prior to the addition of 1 µCi of [3H-methyl]thymidine. The amount of [3H]thymidine incorporated into cellular DNA after 4 h of incubation was determined by liquid scintillation counting. Values represent the mean ± S.D. from five different determinations. **, values differed from no apoE control at p < 0.01; ***, values differed from no apoE PDGF-stimulated control at p < 0.001.


View larger version (15K):
[in this window]
[in a new window]
 
Fig. 3.   Effect of apoE on PDGF-induced proliferation of aortic smooth muscle cells. Serum-starved rat A7r5 smooth muscle cells (panel A) or human coronary artery smooth muscle cells (panel B) were incubated in 96-well plates (5 × 103 cells/well) in DMEM without PDGF and apoE (), or with 10 ng/ml PDGF-BB in the absence (bullet ) or presence (open circle ) of 50 µg/ml apoE. Cell proliferation was determined based on formation of formazan from MTS and phenazine methosulfate as detected by absorbance at 490 nm. Data represent the mean value ± S.D. from five different determinations. *, p < 0.001 versus no apoE control.

ApoE Does Not Affect PDGF Binding to the A7r5 Rat Aortic Smooth Muscle Cells-- The mechanism by which apoE inhibits smooth muscle cell response to PDGF stimulation was investigated initially by determining if apoE interferes with PDGF binding to its receptor on smooth muscle cells. The A7r5 cells were used for these experiments. Results showed that the smooth muscle cells displayed high affinity binding with 125I-labeled PDGF-BB. Excess unlabeled PDGF competitively inhibited 125I-PDGF binding to the A7r5 cells. In contrast, the addition of 50 µg/ml human apoE had no effect on 125I-PDGF binding to the A7r5 cell surface (Fig. 4).


View larger version (16K):
[in this window]
[in a new window]
 
Fig. 4.   Binding of PDGF to A7r5 rat aortic smooth muscle cells. The A7r5 cells were preincubated for 3 h with or without apoE or PDGF prior to the addition of 120 pmol/liter of 125I-PDGF. The incubation was continued for 3 h, the cells were washed, and 125I-PDGF bound to the cells was determined in a gamma  counter. Data represent the mean value ± S.D. from four different determinations. *, p < 0.001 versus control with no apoE or PDGF preincubation.

ApoE Suppression of PDGF-induced Cyclin D1 Gene Expression-- To ascertain the possibility that apoE may interfere with early events in PDGF-induced smooth muscle cell proliferation and migration, experiments were performed to explore the effect of apoE on cyclin D1 gene expression, a prerequisite step for cell transition from the G0 to G1 phase in the cell cycle (22-23). Total RNA was isolated from A7r5 cells incubated with PDGF in the presence or absence of apoE for 6 h, a time period which was reported to result in maximal cyclin D1 mRNA expression after growth factor-induced proliferation of vascular smooth muscle cells (24). Northern blot hybridization of cellular RNA with a rat cyclin D1 cDNA probe revealed a 2.5-fold increase in cyclin D1 mRNA level when cells were challenged with PDGF without apoE (Fig. 5). However, the addition of apoE to the culture media in addition to PDGF reduced the growth factor-induced increase in cyclin D1 mRNA by >50% (Fig. 5).


View larger version (27K):
[in this window]
[in a new window]
 
Fig. 5.   Effect of apoE on PDGF-induced cyclin D1 gene expression in A7r5 rat aortic smooth muscle cells. Quiescent A7r5 cells, plated in 6-well dishes at a density of 1.5 × 105 cells/well, were incubated for 6 h at 37 °C with no stimulation (Control) or stimulation with 10 ng/ml of PDGF in the presence or absence of 50 µg/ml apoE before RNA isolation. Northern blot analysis was performed with 32P-labeled cyclin D1 cDNA and glyceraldehyde-3-phosphate dehydrogenase cDNA probes. The relative amount of cyclin D1 mRNA present in each sample was determined based on the expression level of glyceraldehyde-3-phosphate dehydrogenase mRNA. Data represent average of two independent experiments. The Northern blot data from one experiment is shown in the inset.

ApoE Inhibition of PDGF-induced MAP Kinase in Rat Aortic Smooth Muscle Cells-- An early event important for PDGF stimulation of smooth muscle cells is the activation of MAP kinase (25). Lysate obtained from cells incubated for 5 min with 10 ng/ml of PDGF-BB displayed a 2- to 3-fold increase in MAP kinase activity compared with extracts obtained from unstimulated cells (Fig. 6). However, when 50 µg/ml of human apoE was added simultaneously with PDGF to the cell culture medium, a significant reduction in PDGF-induced MAP kinase activity was observed (p < 0.01, Fig. 6).


View larger version (15K):
[in this window]
[in a new window]
 
Fig. 6.   Mitogen-activated protein kinase activity in A7r5 rat aortic smooth muscle cells. 100 µg of cell lysate, prepared after incubating the A7r5 cells for 5 min at 37 °C with no addition (Control) or with 10 ng/ml PDGF in the presence or absence of 50 µg/ml apoE, was used for the phosphorylation of PHAS-I in vitro. Background phosphorylation was determined by incubating the substrates with buffer in the absence of cell lysate. Data represents the mean value ± S.D. from four different determinations. *, indicates differences at p < 0.01 versus PDGF-stimulated cells.

ApoE Inhibition of oxLDL-Stimulated Smooth Muscle Cell Migration, Proliferation, and MAP Kinase in Rat Aortic Smooth Muscle Cells-- In addition to apoE effects on PDGF-stimulated smooth muscle cell functions, the impact of apoE on oxLDL-stimulated smooth muscle cell response was also explored. Results shown in Fig. 7 indicated that apoE also inhibited the chemoattractive and mitogenic properties of oxLDL on smooth muscle cells (26, 27). This inhibition was also mediated through suppression of the MAP kinase pathway (Fig. 7).


View larger version (17K):
[in this window]
[in a new window]
 
Fig. 7.   Effects of apoE on oxLDL-stimulated smooth muscle cell migration, proliferation, and mitogen-activated protein kinase activity. Smooth muscle cell migration assay (panel A) was determined by incubating quiescent A7r5 cells for 30 min at 37 °C with or without apoE and then added to Transwell membranes in which the lower chamber contained 0.6 ml of DMEM, 0.4% FBS, with or without 200 µg/ml of oxLDL. PDGF stimulation was used as a positive control. Smooth muscle cell proliferation assay (panel B) was determined by incubating quiescent A7r5 cells at 37 °C in DMEM without oxLDL and apoE, or with 5 µg/ml of oxLDL in the absence or presence of 50 µg/ml apoE for 24 h. PDGF stimulation was used as a positive control. MAP kinase activity (panel C) was determined from A7r5 cell lysate prepared after incubating the cells for 5 min at 37 °C with no addition (control) or with 25 µg/ml oxLDL in the presence or absence of apoE. *, p < 0.01 versus stimulated cells.

    DISCUSSION
Top
Abstract
Introduction
Procedures
Results
Discussion
References

Pathological studies indicated that narrowing of the coronary vessels in atherosclerosis and restenosis is related to neointimal hyperplasia, with abnormal proliferation and migration of vascular smooth muscle cells from the tunica media to the intima (28, 29). The current hypothesis suggests that injury to the vascular endothelium exposes the underlying smooth muscle cells to mitogenic growth factors, thereby inducing their phenotypic conversion from the contractile nonproliferating phenotype to that of a secretory proliferating phenotype. Platelet-derived growth factor and oxLDL have been implicated to play a major role in activating both processes of smooth muscle cell migration and proliferation (30-32). The mechanism of cell activation is related to the stimulation of MAP kinase activity (33, 34), which plays critical intermediary roles in mediating signal transduction from the membrane to the nucleus (35). The induction of cell proliferation by MAP kinase has been shown to be a direct result of increased transcription of many immediate early genes (35, 36), including cyclin D1 that is required for cell transition from the G0 to the G1 phase in the cell cycle (37, 38).

A possible role of apoE in modulation of vascular smooth muscle cell growth was suggested by the observation of its synthesis in quiescent but not actively proliferating vascular smooth muscle cells (39). Results described in this manuscript provided the first experimental evidence to document apoE inhibition of agonist-induced smooth muscle cell migration and proliferation. This inhibition was shown to be due to apoE suppression of growth factor-stimulated MAP kinase activity and cyclin D1 gene expression in aortic smooth muscle cells. These in vitro experiments implicated a cytostatic function for apoE in the vessel wall and suggested an additional mechanism by which apoE protects against vascular occlusive diseases.

The results of this study add to a growing list of mechanisms by which apoE is anti-atherogenic. First, apoE reduces hyperlipidemia by mediating cholesterol clearance from circulation (1). Second, apoE suppresses foam cell formation by mediating cholesterol efflux from peripheral cells (40, 41). Third, apoE suppresses lipoprotein oxidation, thereby reducing oxidation-induced endothelial toxicity (42). Fourth, apoE inhibits lymphocyte proliferation (16), thus may limit inflammatory response in the arterial wall. Fifth, apoE inhibits agonist-induced platelet aggregation (43), which will retard the progression of atherosclerotic lesion; and sixth, apoE suppresses growth factor and oxLDL-induced smooth muscle cell migration and proliferation (this study). Taken together, these apoE activities may explain the versatility of this apolipoprotein in conferring protection against various forms of vascular diseases, including cholesterol-induced atherosclerosis (2), angioplasty- or injury-induced restenosis (10, 12, 44), and transplant-induced accelerated arteriosclerosis (46).

In addition to its cell regulatory functions in the cardiovascular system, apoE has also been shown to modulate normal physiology and pathophysiology of other cells and tissues. For example, apoE suppresses steroidogenesis in adrenals and ovaries (47-49). ApoE is also expressed in the brain, where the apoE4 allele is correlated with an increased risk for late onset Alzheimer's disease (50). The relationship between apoE genotype and Alzheimer's disease is related to apoE interaction with beta -amyloid peptide and with tau  in regulation of fibrillogenesis (50, 51). The ability of different apoE isoforms to regulate neurite outgrowth from dorsal root ganglion neurons (52) and to induce neurotoxicity (53, 54) also has been implicated to play a significant role in Alzheimer's disease.

The ability of apoE to modulate such diverse cell functions appears to be related to its ability in modulating cell signal transduction pathways in a cell- and tissue-specific manner. For example, the suppressive effect of apoE on steroidogenesis in adrenals and ovaries is mediated through inhibition of cyclic AMP signaling mechanisms (47-49). In contrast, apoE has no effect on cyclic AMP pathway in lymphocytes, but its inhibition of lymphocyte proliferation was shown to be mediated by suppression of other early cell signaling events that lead to cell arrest at the G1a/G1b boundary (55-57). The mechanism for apoE inhibition of smooth muscle cell proliferation and migration reported herein is different from the apoE effect on lymphocyte proliferation. The smooth muscle cell effect is mediated via inhibition of MAP kinase activity and cyclin D1 activation, resulting in the prevention of cell entry from G0 to G1 phase. Other studies revealed that apoE suppresses agonist-induced platelet aggregation by activation of nitric oxide synthesis (43). ApoE also stimulates nitric-oxide synthase in human macrophages (58). Interestingly, nitric oxide has been shown to inhibit migration and proliferation of vascular smooth muscle cells (59). Whether the effects of apoE on smooth muscle cells reported here are related to its effect on nitric oxide production remains to be determined.

The diverse effects of apoE on cell signaling mechanisms may be related to its ability to bind various different receptors on the cell surface. Currently identified receptors capable of binding apoE include LDL receptor, LRP, gp330/megalin, the VLDL receptor, apoE receptor-2, LR11, and LR7/8 (60, 61). Whereas ligand binding to these receptors usually results in its internalization and degradation in the lysosomes, the potential for signal transduction via ligand interaction with these receptors has not been fully appreciated. There are reports that lactoferrin, lipoprotein lipase, and Pseudomonas exotoxin A binding to LRP on macrophages resulted in a pertussis toxin-sensitive G protein-mediated increase in intracellular calcium ion and inositol triphosphate (62). Significant to the current study is the report that interaction of the 39-kDa receptor-associated protein or anti-LRP with LRP on vascular smooth muscle cells resulted in inhibition of cell migration (63). Whether apoE inhibition of ligand-induced smooth muscle cell migration and proliferation is mediated by LRP, or other members of the LDL receptor gene family present on these cells, such as the VLDL receptor (45), remains to be determined.

    FOOTNOTES

* This research was supported in part by funds from the National Institutes of Health (Grant DK40917) and the Japan Research Foundation for Clinical Pharmacology.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Dagger To whom correspondence should be addressed: Dept. of Pathology and Laboratory Medicine, University of Cincinnati College of Medicine, 231 Bethesda Ave., Cincinnati, OH 45267-0529. Tel.: 513-558-9152; Fax: 513-558-2141; E-mail: Huidy{at}email.uc.edu.

The abbreviations used are: apo, apolipoprotein; VLDL, very low density lipoproteins; IDL, intermediate density lipoproteins; LDL, low density lipoproteins; oxLDL, oxidized low density lipoproteins; PDGF, platelet-derived growth factor; PDGF-BB, homodimer of PDGF B polypeptide; MAP kinase, mitogen-activated protein kinase; DMEM, Dulbecco's modified Eagle's medium; PHAS-I, insulin-regulated phosphorylated heat- and acid-stable protein; FBS, fetal bovine serum; HPF, high power fields; MTS, 3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-4-sulfophenyl)-2H-tetrazolium salt LRP, low density lipoprotein receptor-related protein.
    REFERENCES
Top
Abstract
Introduction
Procedures
Results
Discussion
References

  1. Mahley, R. W. (1988) Science 240, 622-630[Medline] [Order article via Infotrieve]
  2. Shimano, H., Yamada, N., Katsuki, M., Yamamoto, K., Gotoda, T., Harada, K., Shimada, M., and Yazaki, Y. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 1750-1754[Abstract]
  3. Shimano, H., Yamada, N., Katsuki, M., Yamamoto, K., Gotoda, T., Harada, K., Shimada, M., and Yazaki, Y. (1992) J. Clin. Invest. 90, 2084-2091[Medline] [Order article via Infotrieve]
  4. Zhang, S. H., Reddick, R. L., Piedrahita, J. A., and Maeda, N. (1992) Science 258, 468-471[Medline] [Order article via Infotrieve]
  5. Plump, A. S., Smith, J. D., Hayek, T., Aalto-Setala, K., Walsh, A., Verstuyft, J. G., Rubin, E. M., and Breslow, J. L. (1992) Cell 71, 343-353[Medline] [Order article via Infotrieve]
  6. Reddick, R. L., Zhang, S. H., and Maeda, N. (1994) Arterioscler. Thromb. 14, 141-147[Abstract]
  7. Zhang, S. H., Reddick, R. L., Burkey, B., and Maeda, N. (1994) J. Clin. Invest. 94, 937-945[Medline] [Order article via Infotrieve]
  8. Kashyap, V. S., Santamarina-Fojo, S., Brown, D. R., Parrott, C. L., Applebaum-Bowden, D., Meyn, S., Talley, G., Paigen, B., Maeda, N., and Brewer, H. B. (1995) J. Clin. Invest. 96, 1612-1620[Medline] [Order article via Infotrieve]
  9. Shimano, H., Ohsuga, J., Shimada, M., Namba, Y., Gotoda, T., Harada, K., Katsuki, M., Yazaki, Y., and Yamada, N. (1995) J. Clin. Invest. 95, 469-476[Medline] [Order article via Infotrieve]
  10. Seo, H. S., Lombardi, D. M., Polinsky, P., Powell-Braxton, L., Bunting, S., Schwartz, S. M., and Rosenfeld, M. E. (1997) Arterioscler. Thromb. Vasc. Biol. 17, 3593-3601[Abstract/Free Full Text]
  11. Dansky, H. M., Charlton, S. A., Harper, M. M., and Smith, J. D. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 4642-4646[Abstract/Free Full Text]
  12. van Bockxmeer, F. M., Mamotte, C. D. S., Gibbons, F. R., and Taylor, R. R. (1994) Atherosclerosis 110, 195-202[Medline] [Order article via Infotrieve]
  13. Bornfeldt, K., Raines, E., Nakano, T., Graves, L., Krebs, E., and Ross, R. (1994) J. Clin. Invest. 93, 1266-1274[Medline] [Order article via Infotrieve]
  14. Kundra, V., Escobedo, J. A., Kazlauskas, A., Kim, H. K., Rhee, S. G., Williams, L. T., and Zetter, B. R. (1994) Nature 367, 474-476[CrossRef][Medline] [Order article via Infotrieve]
  15. Graf, K., Xi, X. P., Yang, D., Fleck, E., Hsueh, W. A., and Law, R. E. (1997) Hypertension 29, 334-339[Abstract/Free Full Text]
  16. Hui, D. Y., Harmony, J. A. K., Innerarity, T. L., and Mahley, R. W. (1980) J. Biol. Chem. 255, 11775-11781[Abstract/Free Full Text]
  17. Rall, S. C., Weisgraber, K. H., and Mahley, R. W. (1986) Methods Enzymol. 128, 273-287[Medline] [Order article via Infotrieve]
  18. Havel, R. J., Eder, H. A., and Bragdon, J. H. (1955) J. Clin. Invest. 34, 1345-1353[Medline] [Order article via Infotrieve]
  19. Law, R. E., Meehan, W. P., Xi, X. P., Graf, K., Wuthrich, D. A., Coats, W., Faxon, D., and Hsueh, W. A. (1996) J. Clin. Invest. 98, 1897-1905[Abstract/Free Full Text]
  20. Bianchi, S., Fabiani, S., Muratori, M., Arnold, A., Sakaguchi, K., Miki, T., and Brandi, M. L. (1994) Biochem. Biophys. Res. Commun. 204, 691-700[CrossRef][Medline] [Order article via Infotrieve]
  21. Chomczynski, P., and Sacchi, N. (1987) Anal. Biochem. 162, 156-159[CrossRef][Medline] [Order article via Infotrieve]
  22. Won, K. A., Xiong, Y., Beach, D., and Gilman, M. Z. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 9910-9914[Abstract]
  23. Cocks, B. G., Vairo, G., Bodrug, S. E., and Hamilton, J. A. (1992) J. Biol. Chem. 267, 12307-12310[Abstract/Free Full Text]
  24. Fukumoto, S., Nishizawa, Y., Hosoi, M., Koyama, H., Yamakawa, K., Ohno, S., and Morii, H. (1997) J. Biol. Chem. 272, 13816-13822[Abstract/Free Full Text]
  25. Pazin, M. J., and Williams, L. T. (1992) Trends Biochem. Sci. 17, 374-378[CrossRef][Medline] [Order article via Infotrieve]
  26. Autrio, I., Jaakkola, O., Solakivi, T., and Nikkari, T. (1990) FEBS Lett. 277, 247-249[CrossRef][Medline] [Order article via Infotrieve]
  27. Chatterjee, S. (1992) Mol. Cell. Biochem. 111, 143-147[Medline] [Order article via Infotrieve]
  28. Giraldo, A. A., Esposo, O. M., and Meis, J. M. (1985) Arch. Pathol. Lab. Med. 109, 173-175[Medline] [Order article via Infotrieve]
  29. Austin, G. E., Ratliff, M. B., Hollman, J., Tabei, S., and Phillips, D. F. (1985) J. Am. Coll. Cardiol. 6, 369-375[Medline] [Order article via Infotrieve]
  30. Grotendorst, G. R., Chang, T., Seppa, H. E. J., Kleinman, H. K., and Martin, G. R. (1982) J. Cell Physiol. 113, 261-266[Medline] [Order article via Infotrieve]
  31. Jawien, A., Bowen-Pope, D. F., Lindner, V., Schwartz, S. M., and Clowes, A. W. (1992) J. Clin. Invest. 89, 507-511[Medline] [Order article via Infotrieve]
  32. Jackson, C. L., Raines, E., Ross, R., and Reidy, M. A. (1993) Arterioscler. Thromb. 13, 1218-1226[Abstract]
  33. Claesson-Welsh, L. (1994) J. Biol. Chem. 269, 32023-32026[Free Full Text]
  34. Graves, L. M., Bornfeldt, K. E., Sidhu, J. S., Argast, G. M., Raines, E. W., Ross, R., Leslie, C. C., and Krebs, E. G. (1996) J. Biol. Chem. 271, 505-511[Abstract/Free Full Text]
  35. Seth, A., Gonzalez, A., Gupta, S., Raden, D. L., and Davis, R. J. (1992) J. Biol. Chem. 267, 24796-24804[Abstract/Free Full Text]
  36. Chen, R. H., Sarnecki, C., and Blenis, J. (1992) Mol. Cell. Biol. 12, 915-927[Abstract]
  37. Lavoie, J. N., L'Allemain, G., Brunet, A., Muller, R., and Pouyssegur, J. (1996) J. Biol. Chem. 271, 20608-20616[Abstract/Free Full Text]
  38. Abieu, A., Lorca, T., Labbe, J. C., Morin, N., Keyse, S., and Doree, M. (1996) J. Cell Sci. 109, 239-246[Abstract/Free Full Text]
  39. Majack, R. A., Castle, C. K., Goodman, L. V., Weisgraber, K. H., Mahley, R. W., Shooter, E. M., and Gebicke-Haerter, P. J. (1988) J. Cell Biol. 107, 1207-1213[Abstract]
  40. Kruth, H. S., Skarlatos, S. I., Gaynor, P. M., and Gamble, W. (1994) J. Biol. Chem. 269, 24511-24518[Abstract/Free Full Text]
  41. Mazzone, T., and Reardon, C. (1994) J. Lipid Res. 35, 1345-1353[Abstract]
  42. Miyata, M., and Smith, J. D. (1996) Nat. Genet. 14, 55-61[Medline] [Order article via Infotrieve]
  43. Riddell, D. R., Graham, A., and Owen, J. S. (1997) J. Biol. Chem. 272, 89-95[Abstract/Free Full Text]
  44. De Geest, B., Zhao, Z., Collen, D., and Holvoet, P. (1997) Circulation 96, 4349-4356[Abstract/Free Full Text]
  45. Takahashi, S., Kawarabayasi, Y., Nakai, T., Sakai, J., and Yamamoto, T. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 9252-9256[Abstract]
  46. Russell, P. S., Chase, C. M., and Colvin, R. B. (1996) Am. J. Pathol. 149, 91-99[Abstract]
  47. Reyland, M. E., Gwynne, J. T., Forgez, P., Prack, M. M., and Williams, D. L. (1991) Proc. Natl. Acad. Sci. U. S. A. 88, 2375-2379[Abstract]
  48. Dyer, C. A., and Curtiss, L. K. (1988) J. Biol. Chem. 263, 10965-10973[Abstract/Free Full Text]
  49. Reyland, M. E., and Williams, D. L. (1991) J. Biol. Chem. 266, 21099-21104[Abstract/Free Full Text]
  50. Strittmatter, W. J., Saunders, A. M., Schmechel, D., Pericak-Vance, M., Enghild, J., Salvesen, G. S., and Roses, A. D. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 1977-1981[Abstract]
  51. Strittmatter, W. J., Saunders, A. M., Goedert, M., Weisgraber, K. H., Dong, L. M., Jakes, R., Huang, D. Y., Pericak-Vance, M., Schmechel, D., and Rose, A. D. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 11183-11186[Abstract/Free Full Text]
  52. Nathan, B. P., Bellosta, S., Sanan, D. A., Weisgraber, K. H., Mahley, R. W., and Pitas, R. E. (1994) Science 264, 850-852[Medline] [Order article via Infotrieve]
  53. LaFerla, F. M., Troncoso, J. C., Strickland, D. K., Kawas, C. H., and Jay, G. (1997) J. Clin. Invest. 100, 310-320[Abstract/Free Full Text]
  54. Marques, M. A., Tolar, M., and Crutcher, K. A. (1997) Alzheimer's Res. 3, 1-6
  55. Hui, D. Y., Berebitsky, G. L., and Harmony, J. A. K. (1979) J. Biol. Chem. 254, 4666-4673[Medline] [Order article via Infotrieve]
  56. Hui, D. Y., and Harmony, J. A. K. (1980) Biochem. J. 192, 91-98[Medline] [Order article via Infotrieve]
  57. Mistry, M. J., Clay, M. A., Kelly, M. E., Steiner, M. A., and Harmony, J. A. K. (1995) Cellular Immunology 160, 14-23[CrossRef][Medline] [Order article via Infotrieve]
  58. Vitek, M. P., Snell, J., Dawson, H., and Colton, C. A. (1997) Biochem. Biophys. Res. Commun. 240, 391-394[CrossRef][Medline] [Order article via Infotrieve]
  59. Garg, U. C., and Hassid, A. (1989) J. Clin. Invest. 83, 1774-1777[Medline] [Order article via Infotrieve]
  60. Yamamoto, T., and Bujo, H. (1996) Curr. Opin. Lipidol. 7, 298-302[Medline] [Order article via Infotrieve]
  61. Schneider, W. J., Nimpf, J., and Bujo, H. (1997) Curr. Opin. Lipidol. 8, 315-319[Medline] [Order article via Infotrieve]
  62. Misra, U. K., Chu, C. T. C., Gawdi, G., and Pizzo, S. V. (1994) J. Biol. Chem. 269, 18303-18306[Abstract/Free Full Text]
  63. Okada, S. S., Grobmyer, S. R., and Barnathan, E. S. (1996) Arterioscler. Thromb. Vasc. Biol. 16, 1269-1276[Abstract/Free Full Text]


Copyright © 1998 by The American Society for Biochemistry and Molecular Biology, Inc.