Atherosclerosis and Lipoproteins

The Heparin-Binding Proteins Apolipoprotein E and Lipoprotein Lipase Enhance Cellular Proteoglycan Production

Joseph C. Obunike, Sivaram Pillarisetti, Latha Paka, Yuko Kako, Mathew J. Butteri, Yuan-Yaun Ho, William D. Wagner, Nobuhiro Yamada, Theodore Mazzone, Richard J. Deckelbaum, Ira J. Goldberg
https://doi.org/10.1161/01.ATV.20.1.111
Arteriosclerosis, Thrombosis, and Vascular Biology. 2000;20:111-118
Originally published January 1, 2000

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Abstract

Abstract—Apolipoprotein E (apoE) and lipoprotein lipase (LPL), key proteins in the regulation of lipoprotein metabolism, bind with high affinity to heparin and cell-surface heparan sulfate proteoglycan (HSPG). In the present study, we tested whether the expression of apoE or LPL would modulate proteoglycan (PG) metabolism in cells. Two apoE-expressing cells, macrophages and fibroblasts, and LPL-expressing Chinese hamster ovary (CHO) cells were used to study the effect of apoE and LPL on PG production. Cellular PGs were metabolically labeled with 35[S]sulfate for 20 hours, and medium, pericellular PGs, and intracellular PGs were assessed. In all transfected cells, PG levels in the 3 pools increased 1.6- to 3-fold when compared with control cells. Initial PG production was assessed from the time of addition of radiolabeled sulfate; at 1 hour, there was no difference in PG synthesis by apoE-expressing cells when compared with control cells. After 1 hour, apoE-expressing cells had significantly greater production of PGs. Total production assessed with [3H]glucosamine was also increased. This was due to an increase in the length of the glycosaminoglycan chains. To assess whether the increase in PGs was due to a decrease in PG degradation, a pulse-chase experiment was performed. Loss of sulfate-labeled pericellular PGs was similar in apoE and control cells, but more labeled PGs appeared in the medium of the apoE-expressing cells. Addition of exogenous apoE and anti-human apoE antibody to both non–apoE-expressing and apoE-expressing cells did not alter PG production. Moreover, LPL addition did not alter cell-surface PG metabolism. These results show that enhanced gene expression of apoE and LPL increases cellular PG production. We postulate that such changes in vascular PGs can affect the atherogenic potential of arteries.

  • Received September 3, 1998.
  • Accepted June 17, 1999.

ApoE and lipoprotein lipase (LPL) are key proteins in the regulation of lipoprotein metabolism.1 2 3 4 Their actions are interrelated: VLDL and chylomicron removal from the bloodstream requires both of these proteins. LPL hydrolyzes triglyceride in the lipoprotein core and creates a remnant lipoprotein. ApoE mediates clearance of the remnant particles by associating with the LDL receptor and the LDL receptor–related protein on hepatocytes. LPL can also function as an LDL receptor–related protein ligand.5 6 7 Animals that have a gene deletion for apoE have high levels of cholesterol-rich remnants in their bloodstream and extensive lipid-loaded atherosclerotic lesions.8 9 LPL deficiency results in severe hypertriglyceridemia and neonatal demise in mice.10 11

LPL is synthesized primarily in muscle and adipose tissue, and apoE is made in the liver. Both of these proteins are also expressed in several other cell types, including macrophages.12 13 14 Macrophage LPL provides fatty acids for energy.15 Macrophage apoE may facilitate the unloading of cholesterol from cells, thereby promoting the antiatherogenic process of reverse cholesterol transport.16 17 18 19

ApoE and LPL bind to heparin and proteoglycans (PGs) with high affinity. PGs are important constituents of cell membranes and the extracellular matrix20 21 and consist of a core protein to which linear, pendant chains of negatively charged polysaccharides, termed glycosaminoglycans (GAGs), are attached. The 3 major GAG classes in the vessel wall are heparan sulfate (HS), chondroitin sulfate (CS), and dermatan sulfate (DS). PG synthesis requires addition of long GAG chains followed by enzymatic sulfation of these GAGs at different N and O groups. A number of enzymes involved in this process have been cloned.22 23 24 25 26 27 Like PG synthesis, PG catabolism appears to require the participation of several enzymes, including proteases, endoglycosidases, exoglycosidases, and sulfatases.28 29 30 31 32 33 There appears to be an initial, slow internalization of cell-surface PGs followed by a 2-step degradation31 32 33 ; only the second of the 2 steps is inhibited by lysosomal inhibitors. It is unclear whether this process is regulated by ligands that associate with PGs on the cell surface or within the cells.

In previous studies from our laboratory,34 35 we showed that the lipoproteins and LPL that were associated with cell-surface PGs were degraded much more slowly than through the usual receptor-mediated processes. By assessing the rate of turnover of pericellular PGs, we concluded that at least a portion of the LPL and lipoprotein degradation appeared to occur via the uptake and remodeling of cell-surface PGs. Because LPL interaction with PGs is facilitated by low pH,36 we postulated that complexes of PGs and their associated proteins might be more resistant to lysosomal, degradative enzymes. To test this hypothesis, in the present report we studied whether LPL and apoE affect the production of PGs by cells. Because lipoprotein retention within arteries and the actions of growth factors that affect smooth muscle cell proliferation involve vessel wall PGs, we postulated that alterations in the cellular production of PGs by apoE and LPL would modulate atherogenesis.

Methods

Cells

J774 Macrophages

J774 cells were stably transfected to express a wild-type, human apoE cDNA as described previously in detail.18 37 The cells were prepared by cotransfection of the apoE cDNA expression vector containing the human metallothionine IIA promoter with pSV2-neo by calcium phosphate coprecipitation in the presence of 10 mmol/L NH4Cl. This cell line constitutively expresses a human apoE cDNA (E3 isoform). Cells were maintained in 400 μg/mL G418 (Geneticin, Life Technologies) and grown in 10% fetal bovine serum (Gemini Bioproducts Inc) in Dulbecco’s modified Eagle’s medium (DMEM) until the start of the described experimental incubations.

Fibroblasts

ApoE-expressing fibroblasts were derived from F111 rat embryonal fibroblasts.38 Cells were cotransfected with a human apoE cDNA expression vector18 and a pβ-hygro vector conferring hygromycin resistance by calcium phosphate–mediated DNA precipitation.39 Cells were selected, maintained, and grown as mentioned above.

CHO Cells

Chinese hamster ovary (CHO)-K cells were maintained in DMEM/F-12 supplemented with 10% fetal bovine serum. Plasmid pCMV-LPL, in which LPL cDNA was fused to the Rc/CMV vector, was used to transfect CHO cells by using the DNA–calcium phosphate coprecipitation method as previously described.40 LPL-transfected cells were maintained in the same medium with addition of G418. To confirm that the cells were producing LPL, LPL activity was measured before each experiment by using a glycerol triolein emulsion41 as previously described.35

Transfection of the amino-terminal 17% of apoB (apoB17) was performed by using a calcium phosphate–DNA coprecipitate containing 20 g of pCMV5-apoB17-DAF plasmid DNA, 1 g of SV2-neo selection maker, and 20 mol/L chloroquine.42 Cells were selected, maintained, and grown as mentioned above. CHO cells producing apoA1 were generously provided by Dr L. LeCureux (The Upjohn Company, Kalamazoo, Mich).

PG Production

PGs in nontransfected and transfected cells were labeled by incubating them for 20 hours in DMEM containing 10% fetal bovine serum and 50 μCi/mL [35S]sulfate. Cultured medium, pericellular (trypsin-releasable), and intracellular PGs were collected, extensively dialyzed, and assessed in triplicate cultures as previously described.35 Total PG radioactivity was measured after precipitation with 3 volumes of absolute ethanol containing sodium acetate (0.8 g/L) for 18 hours at −20°C. Samples were spun at 2000 rpm for 1 hour, the supernatant was removed, and the pellet was solubilized in 0.5N NaOH. The radioactivity in 1 mL of the aqueous phase was determined by using 3.5 mL of scintillation fluid (Hydrofluor, National Diagnostic) in a model 1800 liquid scintillation counter (Beckman Instruments).

PG Synthesis

Macrophages were labeled with 50 μCi/mL [35S]sulfate for 30 minutes to 4 hours. At the end of each time period, pericellular, intracellular, and secreted PGs were collected and dialyzed extensively. Total PG radioactivity was measured as described above.

Assessment of HSPG and DS/CSPG

Aliquots of dialyzed PGs from the 3 pools were treated with 0.05 U of chondroitinase ABC in enzyme buffer containing 0.01 mol/L N-ethylmalemide, 0.07 mmol/L pepstatin, 0.001 mol/L PMSF (protease inhibitors), 1 mg/mL BSA, and CS (50 μg/mL as a carrier) and then incubated for 20 hours at 37°C. HSPG radioactivity was measured after ethanol precipitation as described above. DS/CSPG was determined by subtracting HSPG from the total PG.

GAG Identification and Size Estimation

For GAG radiolabeling, cells were incubated with medium containing 30 μCi [35S]sulfate and 30 μCi [3H]glucosamine per milliliter at 37°C for 20 hours. PGs from different pools were collected and dialyzed as described above. Protein-free GAG chains were isolated as previously described.43 44 Aliquots (2 mL) of dialyzed PGs were treated with 200 μL of 10N NaOH for 18 hours at 26°C with constant shaking and then neutralized with 10N HCl. To remove core proteins, the samples were adjusted to 7 mol/L urea and loaded onto a column containing 1 mL DEAE cellulose that was washed with 3 bed volumes of 7 mol/L urea, 0.1% Triton X-100, and 0.2 mol/L NaCl in 0.05 mol/L Tris, pH 7.2. 3H-labeled GAGs were eluted with a solution of 7 mol/L urea, 0.1% Triton X-100, and 1 mol/L NaCl in 0.05 mol/L Tris, pH 7.2. Urea was removed by dialysis against 10 mol/L Tris and 0.1% Triton X-100, pH 7.0.

For size analysis, 3H-labeled GAGs from the pericellular pool were chromatographed on a 1.5×90-cm column of Sepharose 6B (Pharmacia) and eluted with 0.2 mol/L NaCl at a flow rate of 11.5 mL/h. Fractions of 0.4 mL were collected and analyzed for radioactivity. Average peak elution position (Kav) of the sample GAG (Ve) was calculated from the equation Kav=(Ve−Vo)/(Vt−Vo), where Vo equals the column void determined by the elution position of LDL, and Vt equals the column total bed volume determined by the elution position of bromophenol blue.45

GAGs of known molecular weights were used to generate a standard curve for molecular weight, and Kav45 molecular weights of the sample GAGs were estimated on the basis of Kav values relative to standard GAGs.

Human Recombinant ApoE

Bacterial recombinant human apoE (apoE3/E3 isoform) was provided by Biotechnology General (Rehovot, Israel) and isolated as previously described.46 This apoE has similar physical and biological properties to native human plasma apoE3.46

Antibodies

Anti-human monoclonal apoE antibody, which interacts with the LDL receptor–binding site of apoE, was kindly provided by Dr Linda Curtiss (The Scripps Research Institute, La Jolla, Calif).

Results

Production of PGs by Transfected Cells

We first examined whether apoE expression by J774 macrophages altered PG production. After incubating the cells for 20 hours in [35S]sulfate-containing medium, the amounts of radiolabel in the medium, pericellular PGs, and intracellular PGs were assessed. As shown in Figure 1A⇓, apoE-expressing cells had 57% more label secreted into the medium pool, 65% more label in the pericellular pool, and 155% more label in the intracellular pool of PGs.

Figure 1.
Figure 1.

ApoE increases cellular PGs. J774 macrophages (A) and fibroblasts (B) were incubated for 20 hours at 37°C in medium containing 50 μCi/mL [35S] sulfate. Medium, pericellular, and intracellular PGs in control cells (closed bar) and apoE-expressing cells (open bar) were assessed after exhaustive dialysis. Data are cpm/3×106 cells and are the mean±SD of experiments performed in triplicate.

To determine whether apoE expression would also enhance PG production by fibroblasts, cells were labeled as mentioned earlier. As shown in Figure 1B⇑, apoE expression by fibroblasts led to an even greater increase in radiolabeled PGs. The apoE-expressing fibroblasts contained ≈174% more PGs in the medium pool, 151% more PGs in the pericellular pool, and 185% more 35S labeled PGs in the intracellular pool. Therefore, apoE expression was associated with a greater amount of sulfate-labeled PGs in both kinds of cells.

Because cells produce several different classes of PGs, PG-degrading enzymes were used to assess the relative increase in HSPG and DS/CSPG. As shown in Figure 2⇓, after 20 hours of labeling, there was a 71% increase in medium HSPG. Similarly, the amount of sulfate label in DS/CSPG was 185% greater in the apoE-producing cells. Therefore, both HSPG and DS/CSPG were secreted in greater amounts from apoE-synthesizing cells.

Figure 2.
Figure 2.

ApoE increases both HSPG and DS/CSPG. Macrophages were labeled with 50 μCi/mL [35S]sulfate for 20 hours at 37°C. Medium PGs were dialyzed and digested with 0.05 U of chondroitinase ABC for 20 hours at 37°C and precipitated with 3 volumes of absolute ethanol containing sodium acetate (0.8 g/L) for 18 hours at −20°C. The precipitates of undigested PGs were regarded as HSPGs. The amount of DS/CSPG was 2-fold higher in macrophages; however, apoE transfection led to increases in both HSPG and DS/CSPG. Medium HSPG and DS/CSPG of control macrophages (closed bar) and of apoE-expressing macrophages (open bar) are shown. Data are cpm/3×106 cells and are the mean±SD of experiments performed in triplicate.

PG Synthesis in ApoE-Transfected Cells

The increase in cellular and pericellular PGs and the increase in secretion of PGs into the medium could have resulted from an increase in synthesis rate or a decrease in PG degradation. To test whether expression of apoE increased PG synthesis, control and apoE-expressing macrophages were incubated with [35S]sulfate for up to 4 hours to assess newly synthesized PGs in different pools. No difference in label incorporation into PGs was noted in the 3 pools during the first hour of incubation (Figure 3⇓). However, later in the incubation, apoE-expressing cells contained more radiolabeled PGs. Similarly, differences in label in the secreted, pericellular, and intracellular PGs were noted beginning at 2 hours, so that at 4 hours the amount of label found in the apoE-expressing cells was 3-fold more in the medium (Figure 3A⇓), 1.6-fold more in the pericellular PGs (Figure 3B⇓), and 2-fold more in the intracellular PGs (Figure 3C⇓). These data suggest that apoE increased either the sulfation or the total amount of GAGs. It should be noted that the labeling procedure of sulfation of GAG chains occurs in the Golgi. This process has different kinetics than that of protein synthesis; this could explain the time required to observe a difference in radiosulfate incorporation.

Figure 3.
Figure 3.

PG synthesis in apoE-transfected cells. Control (closed circle) and apoE-expressing (open circle) macrophages were incubated with 50 μCi/mL [35S]sulfate. Radiolabeled PGs were measured in the medium (A), pericellular (B), and intracellular (C) pools at various times from 0 to 4 hours after exhaustive dialysis. Data are cpm/3×106 cells and are the mean±SD of experiments performed in triplicate.

Comparison of ApoE Effects on GAG Production

The above data show that apoE increased the amount of sulfate label in cellular, pericellular, and medium PGs. To determine whether this was due to sulfation or an increase in the amount or length of the GAG chains, we assessed whether the ratio of glucosamine to sulfate label was similar in control and apoE-expressing cells. For this experiment, apoE-expressing macrophages were doubly labeled with [35S]sulfate and [3H]glucosamine overnight, and the 3 pools of PGs were isolated. The ratio of the 2 labels was the same in the apoE-expressing and control cells (data not shown). Therefore, there was an increase in total GAGs and not a selective increase in sulfation.

To assess whether the increase in GAGs was due to more or longer chains, after the PG core proteins were eliminated, GAG length was measured by elution position on a Sepharose 6B gel filtration column and compared with published calibration curves.45 The gel filtration profile of the pericellular [3H]GAG is shown in Figure 4⇓. Longer GAG chains were found in the pericellular pool from the apoE-expressing macrophages than from non–apoE-expressing cells; the average chain length increased from 7 to 28 kDa. In the first part of the Results, we indicated that the medium PG increase was 174% (Figure 1⇑), so the increase in GAG size can account for probably all of the increase observed in GAG production. Thus, apoE either increased the chain length or protected the GAGs from degradative processes.

Figure 4.
Figure 4.

Elution profiles of [3H]glucosamine-labeled GAGs chromatographed on Sepharose 6B. Control and apoE-expressing macrophages were metabolically labeled with 50 μCi/mL [3H]glucosamine for 20 hours at 37°C. PGs from the pericellular pool were treated with NaOH to remove core proteins and purified by chromatography on DEAE-Sephacel. Protein-free GAGs were subjected to chromatography on Sepharose 6B and eluted with 0.2 mol/L NaCl. Fractions (0.4 mL) were collected and analyzed for radioactivity. Vo (fraction 21) and Vt (fraction 50) were determined by elution position for LDL and bromophenol blue, respectively. GAGs from control cells (closed circle) and from apoE-expressing cells (open circle) are shown.

Effects of ApoE Expression on PG Degradation

One possibility is that the association of apoE protects GAGs from degradation. To study degradation, apoE- and non–apoE-expressing J774 macrophages were labeled for 20 hours, the label was removed, and sulfate-labeled PGs in the 3 pools were assessed during a chase period. Pericellular PG loss was, if anything, more rapid in the apoE-producing cells. As shown in Figure 5⇓, at 30 minutes, 49% of the pericellular sulfate label was lost from the control cells, but 56% of the label was lost from the apoE-expressing cells; at 4 hours, 26% of the original label remained in the pericellular pool of controls and 19% in the pericellular pool of the apoE-expressing cells (Figure 5B⇓). Moreover, as shown previously, there was a marked increase in secretion of PG into the medium. Therefore, it appeared that in the apoE-expressing cells, the larger intracellular and pericellular PG pools led to more PG secretion into the medium (10 890 950 versus 212 502 680 counts per minute per 3×106 cells; Figure 5A⇓). This suggests that apoE increased the amount of GAGs either because longer chains were synthesized or because fewer newly synthesized GAG chains were partially degraded.

Figure 5.
Figure 5.

Effects of apoE expression on PG degradation. Control (closed circle) and apoE-expressing (open circle) macrophages were labeled with 50 μCi/mL [35S]sulfate for 20 hours. The labeling medium was removed, and the cells were washed and incubated for the indicated times in unlabeled medium. After exhaustive dialysis, radiolabeled PGs in the medium (A), pericellular (B), and intracellular (C) pools were assessed. Data are the mean±SD of experiments performed in triplicate.

Effects of LPL Expression on PG Production

Because more PG was produced by apoE-expressing cells, we next tested whether another heparin-binding protein, LPL, would increase cellular PG production. PG production was assessed in control and LPL-overexpressing CHO cells (Figure 6⇓). LPL-transfected CHO cells had 3-fold more sulfate-labeled PGs in cells and in the pericellular pool after a 20-hour incubation. As noted for the apoE-expressing cells, secretion of labeled PGs was much greater in the LPL-expressing cells. PG secretion into the medium at the end of a 20-hour incubation was 185% more in the LPL-expressing cells.

Figure 6.
Figure 6.

LPL increases PG production in CHO cells. Control (closed bar) and LPL-transfected CHO (open bar) cells were incubated for 20 hours at 37°C in medium containing 50 μCi/mL [35S]sulfate. After exhaustive dialysis, medium, pericellular, and intracellular PGs were assessed. Data are the mean±SD of experiments performed in triplicate.

To determine whether the increase in PG production observed in LPL-transfected cells was due to transfection, we assessed PG production in CHO cells transfected with apoA1 and apoB17 (Figure 7⇓). PGs secreted into the medium (Figure 7A⇓), in the pericellular pool (Figure 7B⇓), and in the intracellular pool (Figure 7C⇓) by apoA1- and apoB17-transfected cells were similar to control cells and >1.8- to 2-fold less than LPL-expressing cells. Because an increase in PG production was observed only in LPL-expressing cells but not in apoA1- or apoB17-expressing cells, these data suggest that transfection does not cause increased PG synthesis in cells.

Figure 7.
Figure 7.

Effect of transfection on PG synthesis in CHO cells. Control, LPL, apoA1, and apoB17 CHO cells were incubated for 20 hours at 37°C in medium containing 50 μCi/mL [35S]sulfate. Radiolabeled PGs in the medium (A), pericellular (B), and intracellular (C) fractions were measured after exhaustive dialysis. Data are cpm/3×106 cells and are the mean±SD of 2 experiments performed in triplicate.

Effects of Exogenous ApoE, ApoE Antibody, and LPL on PG Catabolism

We next determined whether the increased GAG secretion into the medium was due to association of the heparin-binding proteins with cell-surface PGs. ApoE-transfected and nontransfected macrophages were incubated with either apoE or LPL, and the turnover of pericellular PGs and accumulation of medium PGs were assessed. The hypothesis was that the binding of these proteins to the cell-surface PGs would protect them from intracellular degradation, so that more of them would be secreted into the medium. Human recombinant apoE (5 μg/mL) with or without 10 μg/mL anti-human apoE antibody was added to control and apoE-expressing macrophages, and PG production after 4 hours was assessed. As expected, apoE-expressing cells produced more PGs (>3-fold) than did non–apoE-expressing cells (Figure 8⇓). However, the addition of neither apoE nor anti-human apoE antibody altered PG production in the medium, pericellular pool, and intracellular pool. Moreover, when apoE-containing lipid emulsion particles were incubated with the cells, PG production was not altered (data not shown). Therefore, it appears that only intracellular production of apoE affected PG production.

Figure 8.
Figure 8.

Effect of exogenous apoE and apoE antibody on PG catabolism. Control (closed bar) and apoE (open bar) macrophages were labeled with 50 μCi/mL [35S]sulfate for 20 hours. The labeling medium was removed, and the cells were incubated for 4 hours in unlabeled medium with or without apoE (5 μg/mL) or anti-human apoE antibody (10 μg/mL). Radiolabeled PGs in the medium (A), pericellular (B), and intracellular (C) pools were measured after exhaustive dialysis. Data shown are cpm/3×106 cells and are the mean±SD of experiments performed in triplicate.

It had been hypothesized that association of proteins with cell-surface PGs would alter PG metabolism.47 Our data concerning the addition of apoE to cells do not appear to support this conclusion. To further test this, LPL, a stronger heparin-binding protein, was added to macrophages. This addition led to no change in the rate of turnover of pericellular, sulfate-labeled PGs Figure 9⇓. Therefore, when LPL was bound to cell-surface PGs, it did not alter their degradation.

Figure 9.
Figure 9.

Effects of LPL on PG catabolism. Macrophages were labeled with 50 μCi/mL [35S]sulfate for 20 hours. The labeling medium was removed, and the cells were incubated for the indicated times in unlabeled medium with or without LPL (10 μg/mL). Pericellular PG turnover was assessed. Data are cpm/3×106 cells and are the mean±SD of experiments performed in triplicate.

Discussion

The present data show that PG-binding proteins modulate PG structure. ApoE expression increased the amount of GAGs associated with PGs in macrophages and fibroblasts; LPL-expressing CHO cells produced more radiolabeled PGs than did control cells. In both cell types, PG levels increased in the cellular, pericellular, and medium pools after the cells were labeled with [35S]sulfate. Similar increases were found when the cells were labeled with [3H]glucosamine. Thus, the amount of carbohydrate in GAGs was increased. Analysis of GAG chain length showed that they were longer in apoE-synthesizing cells.

Why should these PG-binding proteins increase GAG chain length? Either they increased GAG synthesis or decreased its degradation. Data confirming 1 of these processes were not obtained, in part because the complexity of PG assembly complicates analysis of radioisotope labeling studies. Increased synthesis by apoE-producing cells was noted after only 1 hour of labeling. This could have occurred if newly synthesized GAGs were protected from intracellular degradation. Alternatively, the apoE might have bound to nascent GAGs and thereby increased their length.

The enzymes involved in the intracellular PG degradation pathways and their locations are well defined. Studies from other cell types suggest that the initial cleavage of HS GAG is carried out by endoglycosidases that primarily exist in nonlysosomal (chloroquine-insensitive) compartments.31 32 48 The HS intermediates generated by these endoglycosidases are further cleaved, by a chloroquine-sensitive endoglycosidase activity, to smaller oligosaccharides that are rapidly degraded in lysosomes by endoglycosidases. ApoE and LPL might protect GAGs in a chloroquine-sensitive/lysosomal or a low-pH/endosomal compartment, because protein association with GAGs can be greater in more acidic pH. Earlier studies showed that this was true for the LPL association with endothelial cell-surface PG,36 and it was postulated that this was the reason that endothelial cells do not degrade LPL. An alternative to this degradation hypothesis is that apoE and LPL increased the chain length or protected GAGs from the intracellular degradation that occurs during intracellular transport of newly synthesized PGs.

In vitro studies from this and other laboratories have suggested that binding of LPL and basic fibroblast growth factor (bFGF) by HS can protect it from digestion by purified heparanases.49 50 When LPL was preincubated with endothelial HSPG and then subjected to heparanase digestion, LPL protected the fragments of HS from heparanase digestion.49 Other studies showed that the conversion of 85-kDa HS chains to short 6-kDa HS by CHO cell heparanase was inhibited by bFGF.50 A 30- to 60-molar excess of bFGF completely blocked heparanase actions. Our gel filtration data on GAG chain length are consistent with the hypothesis that apoE and LPL blocked GAG degradation. The GAGs in the pericellular pool of apoE-expressing cells were ≈7 to 28 kDa longer than the GAGs from control macrophages. It is conceivable that apoE binding protects GAGs from endogenous GAG-degrading activities.

Perhaps the most important aspect of these studies is the observation that apoE and LPL increased the amount of PG in the medium. There are several pathways responsible for PG delivery to the medium. A pool of newly synthesized PGs is directly secreted from cells. Second, some internalized pericellular PG is delivered into the medium. A third pathway is via release of cell-surface PGs through the actions of extracellular enzymes, a process referred to as “shedding.”32 51 This process may involve degradation of core proteins by proteases or, in the case of glycosylphosphatidylinositol-anchored PGs, by cell-surface, phosphatidylinositol-specific phospholipase C. The protease cleaves the transmembrane core proteins, thus releasing the ectodomains with attached GAGs. Our data are more consistent with apoE’s affecting the first pathway of PG secretion. This is based on the observation that exogenous apoE or LPL, which can associate with pericellular PGs, failed to increase PG levels in the medium. A portion of the newly synthesized PGs in late Golgi vesicles may fuse with endosomes and lysosomes and be degraded, instead of going through the secretory pathway. However, in the presence of bound apoE or LPL, this degradation after fusion may be prevented.

How could apoE protect GAGs in the lysosomal compartment if apoE itself can be degraded by lysosomal proteases? Previous studies have shown that apoE is relatively resistant to intracellular degradation after endocytosis.52 By gel filtration and ultracentrifugation analyses, Chen et al53 recently showed that apoE aggregates at low pH and that this phenomenon may in part be the reason for the insensitivity of apoE to protease degradation. This aggregation can be overcome by increasing the pH.

A final observation from our studies relates to the metabolism of cell-surface PGs to which ligands have been attached. We had previously shown34 that when LPL became associated with the cell surface, some of the uptake and degradation of LPL and its associated lipoproteins appeared to occur exclusive of lipoprotein receptors. Much of this high-capacity but kinetically slower pathway could be accounted for by the turnover of cell-surface PGs; the LPL and lipoproteins were merely bystanders in this process. Because both transmembrane (ie, syndecans) and glycosyl-phosphatidylinositol-anchored PGs are internalized and lead to LPL degradation,35 54 55 there did not appear to be any special role for the transmembrane region of syndecan in this process. Syndecan clusters and is phosphorylated, but the importance of this molecule in ligand and PG metabolism is unknown.51 It has been hypothesized that ligand association with syndecans will cause clustering and thereby alter syndecan metabolism.47 Our studies comparing the turnover of pericellular PGs in the presence and absence of exogenously added LPL do not support this view; LPL association with PGs did not alter PG turnover rates.

A number of PG-mediated processes are thought to be central to the development of the atherosclerotic plaque and may be involved in lipid accumulation, cell proliferation, matrix remodeling, and thrombosis. Both proatherosclerotic and antiatherosclerotic actions of PGs have been described. After crossing the endothelial barrier, apoB-containing lipoproteins may be retained within the arterial wall by their interaction with PGs,56 and smooth muscle cell growth is correlated with the amount of cell-surface PGs.57 Moreover, by sequestering growth factors such as bFGF, matrix PGs prevent growth factor stimulation of cells.58 Matrix metalloproteinases are activated by a PG-bound factor,59 and PG activation of antithrombin and heparin cofactor II60 61 is thought to prevent thrombosis on and within the artery. Finally, PGs in the subendothelial matrix can prevent the association of monocytes62 and Lp(a)63 with matrix adhesive proteins. Thus, depending on the type of PG and the cell of origin, increases in PGs induced by apoE or LPL could modulate pathological responses within the vessel.

In summary, we provide data that 1 process that regulates GAG production by cells is the coproduction of PG-binding proteins. These proteins lead to longer GAG chains. Increases in GAG production and secretion may alter a number of in vivo processes. Because growth factors and cytokines bind to PGs, actions of the former might be modified by altered matrix GAGs. ApoE production by macrophages appears to reduce atherosclerosis in atherosclerosis-prone mice.19 64 65 Perhaps this occurs in part by modulation of the amount or type of GAG that is present within the artery.

Acknowledgments

This work was funded by grants HL-03323 (to J.C.O.), HL-45095 (to I.J.G.), HL-40404 (to R.J.D.), and HL-39653 (to T.M.) from the National Heart, Lung, and Blood Institute, National Institutes of Health, Bethesda, Md. S.P. is an investigator of the American Heart Association, New York City Affiliate. We are indebted to Dr Lyold LeCureux from Upjohn Company (Kalamazoo, Mich) who provided us with apoA1 CHO cells. We wish to thank Tommy Katopodis for maintenance of cell cultures.

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  • The Heparin-Binding Proteins Apolipoprotein E and Lipoprotein Lipase Enhance Cellular Proteoglycan Production
    Joseph C. Obunike, Sivaram Pillarisetti, Latha Paka, Yuko Kako, Mathew J. Butteri, Yuan-Yaun Ho, William D. Wagner, Nobuhiro Yamada, Theodore Mazzone, Richard J. Deckelbaum and Ira J. Goldberg
    Arteriosclerosis, Thrombosis, and Vascular Biology. 2000;20:111-118, originally published January 1, 2000
    https://doi.org/10.1161/01.ATV.20.1.111
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