From the Department of Medicine, Division of
Preventive Medicine and Nutrition, Columbia University College of
Physicians and Surgeons, New York, New York 10032, the
¶ Department of Vascular Biology, Holland Laboratory, American Red
Cross, Rockville, Maryland 20855, and
Smith Kline Beecham
Pharmaceuticals, King of Prussia, Pennsylvania 19406
Received for publication, September 26, 2000, and in revised form, December 11, 2000
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ABSTRACT |
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Lipoprotein lipase (LPL), the major enzyme
responsible for the hydrolysis of circulating lipoprotein triglyceride
molecules, is synthesized in myocytes and adipocytes but functions
while bound to heparan sulfate proteoglycans (HSPGs) on the luminal surface of vascular endothelial cells. This requires transfer of LPL
from the abluminal side to the luminal side of endothelial cells.
Studies were performed to investigate the mechanisms of LPL
transcytosis using cultured monolayers of bovine aortic endothelial cells. We tested whether HSPGs and members of the low density lipoprotein (LDL) receptor superfamily were involved in transfer of LPL
from the basolateral to the apical side of cultured endothelial cells.
Heparinase/heparinitase treatment of the basolateral cell surface or
addition of heparin to the basolateral medium decreased the movement of
LPL. This suggested a requirement for HSPGs. To assess the role of
receptors, we used either receptor-associated protein, the
39-kDa inhibitor of ligand binding to the LDL receptor-related protein
and the very low density lipoprotein (VLDL) receptor, or specific
receptor antibodies. Receptor-associated protein reduced 125I-LPL and LPL activity transfer across the
monolayers. When the basolateral surface of the cells was treated with
antibodies, only anti-VLDL receptor antibodies inhibited transcytosis.
Moreover, overexpression of the VLDL receptor using adenoviral-mediated gene transfer increased LPL transcytosis. Thus, movement of active LPL
across endothelial cells involves both HSPGs and VLDL receptor.
Lipoprotein lipase
(LPL)1 is a 120-kDa dimeric
protein that associates with the luminal surface of endothelial cells
in multiple organs but especially in cardiac and skeletal muscle and in
adipose tissue (1). This enzyme hydrolyzes the triglyceride in
circulating lipoproteins such as chylomicrons and VLDL and produces
free fatty acids that are used for metabolic energy or for fat storage.
Endothelial cells do not synthesize LPL; rather myocytes and adipocytes
produce it. Thus, it is a protein that requires transcytosis across the endothelial cell barrier, in this case from the interstitial fluid to
the luminal side of the cells.
There are several possible ways that LPL could cross the endothelial
barrier. Nonspecific transport of molecules across endothelial monolayers occurs either via paracellular routes between the cells or
via vesicular transit through cells (2). Alternatively, a specific
transcytosis pathway could exist which requires LPL to associate with a
cell surface receptor and then transports LPL through the cells. This
process would be analogous to that which transfers IgA across
epithelial cells (3). The first step in a specific LPL transcytosis
pathway would involve LPL interaction with the basolateral side of
endothelial cells. LPL binds to a number of cell surface molecules
including heparan sulfate proteoglycans (HSPGs) and members of the LDL
receptor family (4). In bovine endothelial cells the most highly
expressed of these receptors is the VLDL receptor (VLDLr) (5). A
previous study suggested that HSPGs are required for LPL transcytosis
(6). It is, however, unclear whether HSPGs are sufficient for transport or whether HSPGs must operate in concert with receptors. The binding of
LPL to several members of the LDL receptor family leads to uptake and
degradation of LPL by cells. There are no data on whether these
receptors participate in transendothelial movement of LPL or other ligands.
In this report, we present data showing that LPL transcytosis across
endothelial monolayers requires both HSPGs and the VLDLr. LPL
transcytosis was diminished by removal of HSPGs and inhibition of
receptors by RAP, a 39-kDa protein that was copurified with the LDL
receptor-related protein (LRP) (7). This protein binds to members of
the LDL receptor family and inhibits ligand binding and uptake by those
receptors (8, 9). Furthermore, antibodies against the VLDLr blocked LPL
translocation and increased expression of this receptor-increased
transcytosis. Thus, LPL requires both HSPGs and receptors for
translocation across endothelial cells.
Purification and Radioiodination of LPL--
LPL was purified
from unpasteurized bovine milk according to the method of Socorro
et al. (10) with modifications as described by Saxena
et al. (11). 300-500 µg/ml purified enzyme was stored at
LPL was radioiodinated enzymatically with glucose oxidase and
lactoperoxidase (12). Radioiodinated LPL was purified by
heparin-agarose (Bio-Rad) affinity chromatography and stored at
Endothelial Cell Monolayers--
Primary cultures of bovine
aortic endothelial cells (BAECs) were established as reported (13) and
were grown in DMEM containing 10% fetal bovine serum (Gemini
Bioproducts Inc., Calabasas, CA), 1% (v/v) penicillin and streptomycin
solution, and 1% (v/v) glutamine solution (both from Life
Technologies, Inc.). Polarized BAEC monolayers were grown on gelatin
and fibronectin-coated polyethylene terephthalate 10-mm filters (pore
diameter, 3.0 µm) (Becton Dickinson Labware, Franklin Lakes, NJ).
This allowed access to both the basolateral side of the cells adjacent
to the lower chamber and the apical cell surface in contact with medium
in the upper chamber (14). Approximately 4-5 × 104
cells were seeded onto the filters. Experiments on nonviral infected cells were conducted 5-6 days after seeding the endothelial cells. The
media in the upper chamber (0.5 ml) and lower chamber (1 ml) were
changed every other day. Movement of both [3H]dextran (70 kDa, American Radiolabeled Chemicals, St. Louis, MO) and LDL was
routinely assessed to verify that the cell monolayer was intact.
125I-LPL Transport across
Monolayers--
RAP-sensitive transport of 125I-LPL across
the monolayers was studied after adding 1 µg/ml radiolabeled LPL to
DMEM and 1.5% BSA in the basolateral chambers. In some experiments,
the basolateral side of the cells was incubated with 2.5 units/ml
heparinase/heparitinase (Sekagaki America Inc., Bethesda, MD) or 5 µg/ml RAP-containing medium. After 1 h, the medium was removed,
and the cells were washed prior to adding 125I-LPL.
125I-LPL that appeared in the upper chamber was monitored
over time by removing 100 µl of medium from the apical side of the
cells at 30, 60, 120, and 180 min. The chambers were not stirred to avoid disruption of the monolayers. All experiments were performed using triplicate chambers. At the conclusion of the experiments, the
cells were washed, and 125I-LPL associated with luminal and
basolateral surfaces was released by the addition of DMEM-BSA
containing 100 units/ml heparin (Elkins Sinn, Cherry Hill, NJ) at
4 °C for 30 min to upper and lower chambers. Intracellular
125I-LPL was then estimated by dissolving the
heparin-treated cells in 0.1 N NaOH and by measuring the
radioactivity. In other experiments, heparin was added to the
basolateral chamber along with the 125I-LPL, and the
appearance of 125I-LPL protein in the upper chamber was
determined. The radioactive protein was routinely immunoprecipitated
with 10% trichloroacetic acid; in all experiments less than 10% of
the counts in the chambers were not precipitated
Transport of LPL Activity across Endothelial Cell
Monolayers--
To study the transport of LPL activity, 100 µg/ml
LPL purified from bovine milk as described earlier was added to the
basolateral medium under different conditions. Because LPL rapidly
loses its activity during a 37 °C incubation and is less stable in
more dilute solutions, this higher concentration of LPL was required to
measure transport of its activity. In addition, the activity was
measured at an earlier time (1-2 h). In some experiments, LPL was
added together with 5 µg/ml RAP or 5 units/ml heparin or after a 1-h
incubation of the cells with RAP and/or VLDLr antibodies. 100 µl of
medium from the upper chamber was collected at 60 and 120 min and
immediately frozen at Effects of RAP and Antibodies on LPL Transport--
RAP was
produced as a fusion protein with glutathione S-transferase
in an expression system utilizing human placental RAP cDNA (7).
Antibodies against RAP (Rb80), LPL and blocking antibodies against the
LDL receptor, VLDLr, and LRP were described previously (16-19).
Adenovirus Expression of VLDLr--
For expression of VLDLr in
cells, BAECs were infected with adenovirus-containing human VLDL
receptor (AdhVLDLr) and Ligand Blotting Analyses--
Cell extracts from control,
AdLacZ-, and AdhVLDLr-infected BAECs were prepared as described (20).
Cells on 10-mm filters were dissolved in 0.05 ml of ice-cold solution
containing 50 mM HEPES, pH 7.4, 0.5 M NaCl,
0.05% Tween 20, 1% Triton X-100, 1 mM phenylmethysulfonyl
fluoride, 25 µg/ml leupeptin, and 2 µg/ml pepstatin. The cell
extract was sheared with a 21-gauge needle and then centrifuged at
14,000 rpm for 10 min. The supernatant of each condition in triplicate
was collected and pooled. For RAP ligand blotting, an aliquot (10 µl)
was run on 5% SDS-polyacrylamide gel electrophoresis under nonreducing
conditions and electrophoretically transferred to a nitrocellulose
membrane. The membrane was incubated with 25 nM RAP in
phosphate-buffered saline containing 3% nonfat milk, 0.05% Tween 20, and 5 mM CaCl2 for 1 h at 25 °C
(blocking buffer). The membrane was then incubated with anti-RAP IgG
(Rb80, 1 µg/ml) in blocking buffer for 1 h at 25 °C and
washed three times in phosphate-buffered saline containing 0.1% Tween
20. The membrane was then incubated with a donkey anti-rabbit IgG
horseradish peroxidase conjugate (Bio-Rad) for 1 h at 25 °C.
After washing, the bands were visualized using the ECL kit (Amersham
Pharmacia Biotech).
Transport of 125I-LPL across Endothelial Cell
Monolayers--
We first assessed how increasing amounts of LPL added
to the basolateral side of BAECs affected the amount of LPL that
crossed the monolayer. As shown in Fig.
1A, increasing
125I-LPL in the medium on the basolateral side of the cells
led to more 125I-LPL in the upper chamber. However, when
the 125I-LPL concentration exceeded 1 µg/ml, a lower
percentage of the LPL was transported. For that reason, 1 µg/ml was
used for subsequent experiments.
To determine the specificity of LPL transport, we used unlabeled LPL to
inhibit 125I-LPL transport. The addition of 300 µg/ml LPL
to the lower chamber in the presence of 1 µg/ml 125I-LPL
decreased the appearance of 125I-LPL in the upper chamber
to 46 ± 5% of control (Fig. 1B). The amounts of
125I-LPL within the cells and released from the apical
surface are shown in Fig. 1, C and D. More
125I-LPL was within the cells than on the cell surface.
Increasing amounts of 125I-LPL in the lower chamber led to
more labeled LPL in the cells and on the apical surface. In the
presence of an excess of unlabeled LPL, both the intracellular and cell
surface-labeled LPL were decreased by >50%. Thus cellular uptake and
transfer to the apical surface were inhibited, but not completely.
Effects of Modulation of HSPGs on LPL Transport--
Although the
role of HSPGs and members of the LDL receptor family as LPL-binding
molecules is well documented, only limited studies (22, 23) have been
performed with respect to their role as transcytosis molecules. Because
members of the this family of receptors often act in concert with HSPGs
(22, 23), we tested whether LPL movement across monolayers requires
association with HSPGs. 125I-LPL translocation across BAEC
monolayers was studied in the presence of heparin. As shown in Fig.
2, heparin at 5 units/ml decreased
125I-LPL in the upper chamber after 3 h by >54%.
Inhibition also was found with a higher dose of heparin (50 units/ml).
To test whether HSGP degradation affected LPL transfer, HSPGs on the
basolateral side of the cells were removed by incubating the cells with
HSPG-degrading enzymes. The results shown in Fig. 2, inset,
demonstrate that heparinase/heparinitase treatment of the basolateral
side of the cells reduced 125I-LPL movement across the
monolayers, a 41% inhibition. Thus, optimal LPL movement across the
monolayers required its association with proteoglycans on the
basolateral side of the endothelial cells. However, a large amount of
the radiolabeled LPL was not affected by unlabeled LPL, heparin, and
heparinase. This suggested that the radiolabeled LPL was tracing two
different transport pathways only one of which was reduced by unlabeled
LPL and inhibition of LPL-HSPG interaction.
Effects of RAP on LPL Transport across Endothelial Cell
Monolayers--
We next determined whether HSPGs are sufficient for
LPL transcytosis or whether they serve as accessory molecules for
transcytosis receptors. To test whether LRP, VLDLr, or other members of
this family are involved in LPL transport, 1 µg of
125I-LPL was added to the lower chamber of monolayers that
had been incubated for 1 h with RAP. The RAP had been added to the
lower chamber in various concentrations, 0.33-10 µg/ml, to produce a molar ratio of 1:1-1:30 of LPL to RAP assuming that LPL is an ~120-kDa dimer. In a dose-dependent manner, RAP decreased
125I-LPL movement from the lower to the upper chamber (Fig.
3A). The highest doses 5 µg/ml (shown as filled squares) and 10 µg/ml (shown as open triangles) reduced the amount of
125I-LPL >40%; 3.3 µg (shown as filled
circles) reduced it >30%. The maximum reduction of
125I-LPL transport with 5 µg/ml was 40-80% depending on
the experiment. Moreover, the percentage of inhibition appeared greater
at 2 than at 3 h; >50% inhibition was found using the two
highest concentrations of RAP.
We next tested whether the RAP- and heparin-mediated inhibition were
additive. As shown in Fig. 3B, when heparin was added to the
RAP-treated cells the inhibition of LPL transport was similar to that
found using only RAP. The addition of excess unlabeled LPL also did not
lead to more inhibition of transport than that found with RAP alone.
This suggests that the three interventions, RAP, excess unlabeled LPL,
and heparin, were inhibiting the same pathway.
To assess whether our interventions affected movement of a control
molecule, 1 µg/ml dextran (molecular weight 70,000) was added to the
basolateral side of the cells, and its movement over the ensuing 3 h was assessed. Dextran movement to the upper chamber is shown in Fig.
3B; this was not affected by heparin or the addition of
RAP.
Movement of LPL from the apical to the basolateral side of the
monolayers was also studied in the presence and absence of RAP and
using cells that were treated with heparinase/heparitinase. LPL
movement was much greater in this direction (Fig. 3D,
compare the total transport across the cells with that in Fig.
3A). As had been found for movement in the opposite
direction, both RAP and removal of glycosaminoglycan chains from HSPGs
decreased the amount of 125I-LPL appearing in the medium on
the basolateral side of the cells.
Effects of RAP and Heparin on Cell Surface and Intracellular
125I-LPL--
The amount of radioactive LPL within the
cells and on the apical surface was determined after a 3-h
incubation ± RAP (Fig. 4). RAP
reduced the amount of heparin-releasable 125I-LPL >50%.
In the experiment illustrated in the figure that equaled a reduction
from 1.05 ± 0.07 to 0.52 ± 0.12 ng/cell protein.
Intracellular 125I-LPL was reduced from 11.68 ± 1.98 to 6.75 ± 0.99 ng/cell protein by RAP. Treating the cell
monolayers with heparin alone, or heparin plus RAP, also decreased both
cell surface and intracellular LPL. Heparin was more effective than
RAP, possibly because it decreased both receptor and nonreceptor
HSPG-mediated uptake of 125I-LPL. Adding RAP to heparin led
to no further decrease in intracellular or cell surface
125I-LPL. Because RAP and heparin decreased both LPL in the
upper chamber and in the cells, it suggested that the LPL movement was via an intracellular process.
Effects of Heparin and RAP on LPL Activity Transport across
Endothelial Cell Monolayers--
To investigate the effect of RAP and
heparin on the appearance of LPL activity on the apical side of the
endothelial cell monolayer, 100 µg/ml purified LPL was added to the
lower chamber in the absence or presence of 5 units/ml heparin, 5 µg/ml RAP, or to monolayers in which the basolateral side of the
cells was treated with 5 µg/ml RAP. As shown in Fig.
5, treatment of the cells with RAP
(denoted RAP treated) and addition of RAP along with LPL
(denoted LPL+RAP) or adding heparin (denoted
Heparin) reduced the amount of LPL activity appearing in the
upper chamber. At 1 h, all of the treatments decreased LPL
activity transport by >90% compared with control cells. A similar
inhibition of LPL activity movement through the endothelial monolayer
was found at 2 h (data not shown). Therefore, RAP and heparin
markedly decreased the amount of LPL activity transferring from the
lower to the upper chamber. This effect of RAP and heparin on LPL
activity transcytosis was much greater than that on
125I-LPL transport.
Transcytosis of Heat-inactivated LPL--
One possible reason for
the greater effect of RAP on LPL activity than 125I-LPL was
that inactive LPL protein was transported across the monolayers by a
non-RAP-inhibited process. It should be noted that during these
experiments, some iodinated LPL would have been converted to inactive
monomer, and the data using this tracer would assess both active and
inactive LPL. To determine whether inactivated LPL was transported in a
manner similar to that of active dimeric LPL, heat-inactivated
125I-LPL was studied. These preparations have been
characterized previously and consist primarily of inactive LPL that
elutes from heparin at a lower salt concentration and is thought to be
monomeric (23). Our preparation was assessed by SDS-polyacrylamide gel electrophoresis and consisted primarily of an ~55-kDa protein; however, ~ 80% of this preparation eluted from heparin affinity gel
with 0.5 M NaCl-containing buffer. This is in contrast to nonheated LPL, in which <20% of the 125I-LPL eluted at
this salt concentration. As shown in Fig.
6, heat-inactivated 125I-LPL
(denoted Inactive LPL) was transferred from the lower to the
upper chamber nearly twice as fast as nonheated 125I-LPL
(denoted Active LPL). Moreover, this transfer was unaffected by either the addition of RAP along with the LPL, or preincubation of
the endothelial cells with RAP. Therefore inactive 125I-LPL
crosses the monolayer at a greater rate via a non-RAP-sensitive pathway.
Effects of Antibodies to LPL and Receptors--
Antibodies (IgG,
30 µg) against LPL or members of the LDL receptor family were added
to the lower chamber prior to the addition of 125I-LPL.
These amounts of antibodies were sufficient to inhibit the respective
receptors (24) or to bind to each LPL molecule. As shown in Fig.
7A, anti-VLDLr and anti-LPL
antibodies inhibited transcytosis by ~50%; these data are shown in
the open squares and open inverted
triangles, respectively. In contrast, antibodies to the LDL
receptor (open circles) and LRP (filled
triangles) had no effect. The same antibody treatment was used to
assess the role of the VLDLr in transport of active LPL (Fig.
7B). Anti-VLDLr antibodies decreased LPL movement to the
apical side of the cells by >80%. This inhibition was comparable to
that found with RAP; RAP and anti-VLDLr antibody did not have an
additive effect on inhibition of transport. Therefore, inhibition of
the VLDLr decreased LPL transport across the cultured endothelial
cells.
VLDLr Overexpression Increases LPL Transport across Endothelial
Cell Monolayers--
We next tested whether more VLDLr expression
increases LPL transcytosis. BAECs were infected with either AdhVLDLr or
AdLacZ, and the expressed VLDLr was examined by RAP ligand blotting of membrane extracts (Fig. 8A).
In control and infected cells a strong band for a protein of
Mr~120,000 which corresponded to the VLDLr was
found. The identity of this band was confirmed using anti-VLDLr antibodies. Only AdhVLDLr-infected cells had an intensely staining human VLDLr band, seen in Fig. 8A, lane 3. BAECs
express a lower molecular weight form of the VLDLr than human cells
(5). Two other less intense high molecular weight bands were observed. The second band reacted with anti-LRP antibodies; the highest molecular
weight protein was, presumably, megalin.
As expected, VLDLr expression increased LPL transport. A time course of
125I-LPL transcytosis across control and infected BAECs is
shown in Fig. 8B. Cells infected with AdhVLDLr are shown in
filled circles, control cells are shown in open
squares, and LacZ-expressing cells are indicated by
filled triangles. VLDLr overexpression increased 125I-LPL transport > 60% in BAECs after 180 min. RAP
completely blocked the increased transcytosis due to AdhVLDLr infection
and reduced the transport to below that of control cells. These data
confirm that LPL transport across endothelial cells can be mediated by the VLDLr.
Fig. 8, C and D, shows the effects of VLDLr
overexpression on apical cell surface and intracellular
125I-LPL, respectively. BAECs infected with AdhVLDLr had
more intracellular and more cell surface 125I-LPL than
control cells; 125I-LPL on the apical surface was
approximately doubled, and that inside the cells was increased >60%.
This effect was completely blocked by RAP. Thus, VLDLr overexpression
increased cellular uptake and transfer of LPL across the monolayer.
Our experiments demonstrate that LPL transport from the
basolateral to the apical side of cultured endothelial cell monolayers is modulated by two processes: LPL binding to HSPGs and interaction with the VLDLr. Our previous (6) and current experiments confirm the
participation of HSPGs in the LPL transcytosis process. As has been
shown for endocytosis, our experiments suggest that the role of HSPG
binding is to concentrate the LPL on the cell surface and increase the
efficiency of its binding to cell surface receptors.
Our data suggested that receptors, especially the VLDLr, were involved
in LPL movement across the monolayers. 1) Incubation of the monolayers
with RAP decreased the transfer of radioactive LPL from the abluminal
to luminal side of the cells. 2) Transfer of LPL activity to the apical
side of the cells was inhibited to an even greater degree by RAP. 3)
Antibodies against the VLDLr decreased LPL transfer. 4) Overexpression
of the VLDLr in endothelial cells increased LPL movement.
Excess of unlabeled LPL markedly reduced 125I-LPL transport
across BAEC monolayers. However, we noted that the level of competition did not exceed 50%. Furthermore, when heparin, RAP, or VLDLr
antibodies were used to block 125I-LPL transcytosis, the
effect was no greater than 60% in most of the experiments. Important
for the interpretation of these experiments is the observation that
purified LPL rapidly converts to an inactive monomeric form during a
37 °C incubation (26). For that reason, our experiments using
125I-LPL assessed the transfer of both active and inactive
LPL. Although this complicated both the experiments and interpretation,
we were still able to define cellular requirements needed for this
process. Two types of experiments clarified this situation. Studies
were performed using heat-inactivated 125I-LPL, and others
assayed movement of LPL activity. Heat-inactivated LPL transferred
across the monolayer more rapidly, and its transport was not decreased
by RAP. In contrast, LPL activity transcytosis was almost completely
blocked by heparin, RAP, or VLDLr antibodies. Therefore, monomeric and
active dimeric LPL differed in their transport. Active LPL transcytosis
occurs via a HSPG-requiring and RAP-sensitive pathway, whereas
presumably smaller inactive 125I-LPL monomers transfer more
quickly through nonspecific paracellular or transcellular routes.
Both HSPGs and the VLDLr modulated intracellular and cell surface LPL.
Much more 125I-LPL was found inside than on the apical
surface of the cells. This was not surprising because endothelial cells
internalize and recycle LPL (25), and some of the intracellular LPL may have been destined for recycling. All of the interventions that decreased the amount of 125I-LPL in the upper chamber
medium also decreased apical cell surface and intracellular
125I-LPL: heparin, heparinase, excess unlabeled LPL, RAP, and
anti-VLDLr antibodies. Therefore, movement of LPL to both the surface
and into the medium is likely to be via the same process. We postulate that internalized LPL translocates across the cell only if it is
targeted via the initial VLDLr uptake.
Endothelial cells, unlike other cells, degrade very little internalized
LPL (25). A similar observation was noted by Friedman et al.
(27) and Argraves et al. (20). We tested the BAECs used in
these experiments and found that like porcine cells (32) <5% of cell
surface 125I-LPL was degraded by the cells. Although the
VLDLr mediates LPL degradation in other cells, receptors in this family
do not always lead to degradation of internalized ligands. It has been
shown recently that hepatic uptake of apoE leads to recycling of this apoprotein (28). Like LPL, apoE is a strong heparin-binding protein
that also interacts with LRP. Thus, it is likely that the cell type
determines whether these receptors participate in protein transcytosis,
retroendocytosis, or degradation. In agreement with this hypothesis, it
has been reported recently that gp330 is responsible for transcytosis
of thyroglobulin by thyroid cells grown on filters (29) even though
this receptor usually leads to lysosomal degradation of ligands.
IL-8 is another protein that binds to both proteoglycans and cellular
receptors (30, 31). Unlike LPL, this protein enters the bloodstream via
the postcapillary venules and then circulates in the blood. A
histological study (32) showed that injection of heparanase blocked
IL-8 transcytosis, i.e. the IL-8 remained in the injected
tissues presumably because it was unable to bind to proteoglycans, a
step required for interaction with the IL-8 receptor. Our current
studies suggest that a similar process is required for LPL. Although
our observations in cultured BAECs implicate HSPGs and VLDLr as
components of a LPL transport system one must question whether these
observations are of physiologic importance for LPL actions. Muscle and
adipose tissue are the two most important sites of LPL-mediated
hydrolysis of lipoprotein triglyceride. These are also sites of
abundant VLDLr expression (17). This receptor is highly expressed in
endothelial cells (33) and, therefore, is in the right tissues and
cells to transport LPL. Moreover, the VLDLr is modulated by feeding and
fasting, and its expression parallels changes in LPL activity in those tissues (34).
The observation that RAP inhibits the transcytosis of LPL to its site
of action is consistent with and explains several in vivo
observations. Overexpression of RAP in mice consistently increased
circulating triglyceride levels; increased numbers of remnant
lipoproteins were found only in LDL receptor knockout mice (35).
RAP-induced hypertriglyceridemia was associated with an increase in the
size of circulating triglyceride-containing lipoproteins (36). In mice
lacking LRP in the liver, RAP overexpression reduced postheparin LPL
activity (21). Therefore, RAP has an action other than inhibiting
chylomicron remnant uptake via liver LRP. In experiments not shown we
have found that RAP neither inhibits LPL binding to the luminal side of
endothelial cells nor inhibits LPL-mediated hydrolysis of emulsion or
VLDL triglyceride, an observation also made by others (21). We propose
that inhibition of LPL transcytosis is the cause of RAP-induced hypertriglyceridemia.
One seemingly inconsistent piece of data is the lack of an abnormal
lipoprotein phenotype in VLDLr knockout mice (37). These mice, however,
are less obese than their littermates. Of note, mice not expressing LPL
in fat also have a normal lipoprotein phenotype, but when crossed
with ob/ob mice they are less obese (38). One possible explanation for
the lack of hypertriglyceridemia in the VLDLr knockout mice is that LPL
transcytosis is not rate-limiting. It should be noted that postheparin
plasma LPL activity in mice is ~5 times greater than in humans (39),
therefore a fraction of normal LPL activity may be sufficient for
normal lipolysis. In support of this hypothesis is a recent report
showing that mice deficient in both VLDLr and the LDL receptor develop
hypertriglyceridemia when they are placed on a high fat diet (40).
Thus, when mice have greater lipoprotein flux through the circulation
lipolysis may be limited by VLDLr deficiency.
In previous studies from this laboratory we studied LPL interaction
with the surface of endothelial cells. With a number of colleagues, we
reported that LPL binds to a 200-kDa transmembrane heparan sulfate
proteoglycan on the luminal surface of endothelial cells (41). This
cell surface proteoglycan is syndecan 4. LPL binding is mediated by the
glycosaminoglycan chains on the proteoglycans, and a highly sulfated
decasaccharide is required for optimal LPL binding (42). We also
reported that LPL bound to a 116-kDa protein that was not a
proteoglycan (43). A protein was isolated and sequenced and was an
amino-terminal fragment of apolipoprotein B (44); this portion of apoB
associates with LPL and heparin (45). The VLDLr has a molecular
mass of 120 kDa (5). It is likely that our attempts to identify
LPL-binding proteins by ligand blotting and LPL affinity chromatography
missed the VLDLr because of its size similarity to the more abundant
apoB fragment.
In summary, we have shown that LPL transcytosis across endothelial
cells is inhibited by reducing LPL interaction with HSPGs and by
blocking the VLDLr. Overexpression of the VLDLr increases this process.
In addition, a second pathway that allows for movement of inactive LPL
molecules also exists and is not sensitive to RAP or heparin. These
data suggest that HSPGs participate with members of the LDL receptor
family to internalize and transfer ligands from the interstitial space
to the circulation. This process may be important for the extracellular
transport pathway of active LPL. Similar receptor-mediated pathways are
likely to be involved in the physiological functioning of a number of
other proteins that must transfer from parenchymal cells to the circulation.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
70 °C. Enzyme activity was assayed with a glycerol-containing triolein emulsion as described previously (11). The purified enzyme had
a specific activity of 40-50 mmol of oleic acid released/h/mg of
enzyme at 37 °C.
70 °C. Typical specific activity of the preparation was
1,000-2,000 cpm/ng, and >90% of the radioactivity was precipitated
with trichloroacetic acid. 125I-LPL was purified by
Sephadex G-25 gel filtration (PD-10, Amersham Pharmacia Biotech) prior
to use to remove degradation products. Heat-inactivated LPL was
prepared by heating LPL for 1 h at 52 °C.
70 °C. The samples were subsequently defrosted and assayed together. For these measurements we used a more
sensitive LPL activity assay described by Hocquette et al.
(15), and all assays were performed in triplicate. 20% Intralipid (Amersham Pharmacia Biotech) was diluted with an equal amount of
deionized water to produce a 10% soybean oil emulsion. To incorporate the radiolabel, the emulsion was sonicated (75 W, 10 min, 50% pulse
mode) with 378 µCi of [3H]triolein (specific activity,
21 Ci/mmol, Amersham Pharmacia Biotech). 30 µl of cell culture medium
was incubated for 1 h at 25 °C with 10 µl of the Intralipid
emulsion, 10 µl of heat-inactivated serum as a source of
apolipoprotein C-II, 100 µl of incubation buffer (0.2 M
NaCl, 0.3 M Tris-HCl, pH 8.5, 6% fatty acid-free bovine
serum albumin, Sigma fraction V), 2 units of heparin/ml, and 50 µl of
deionized water. The reaction was terminated by the addition of 0.5 ml
of deionized water and 2 ml of a solution containing isopropyl
alcohol/heptane/H2SO4 (48:48.3:1 v/v/v).
After a 3-min centrifugation at 3,000 × g 800 µl of
the upper phase was transferred to new tubes containing 3 ml of heptane
and 1 ml of of alkaline ethanol, water, 2 M NaOH
(500:475:25 v/v/v). A second centrifugation was performed as above, and
the upper hydrophobic phase was replaced by 3 ml of heptane. The
samples were then centrifuged again in the same manner. The
radioactivity in 800 µl of the remaining basic hydrophilic phase was
determined using 3.5 ml of scintillation fluid (Ecoscint H, National
Diagnostic, Atlanta, GA) in a model 1800 liquid scintillation counter
(Beckman Instruments).
-galactosidase-expressing adenovirus
(AdLacZ) when the cells were 80-90% confluent; experiments using
these cells were conducted 24 h after infection. The barrier function of the endothelial cell monolayers was examined using trypan
blue and dextran transport as described previously (6).
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Fig. 1.
Effects of different doses of LPL on
transport, medium, and intracellular LPL. Panel A, LPL
transport. Confluent BAEC monolayers grown on semipermeable membranes
were incubated for 3 h with the indicated amounts of
125I-LPL in the medium on the basolateral side of the
cells. The amount of radioactivity appearing in the upper chamber was
measured over time. The radioactivity in the medium was precipitated
with 10% trichloroacetic acid; >90% of the radioactivity was
precipitated. Shown are averages ± S.D. of experiments performed
in triplicate. Panel B, effect of adding unlabeled LPL. In a
separate experiment performed as described in panel A, 300 µg/ml unlabeled LPL was included in the medium on the basolateral
side of the endothelial monolayer along with 1 µg/ml
125I-LPL. The percentage of radioactive LPL transferred to
the upper chamber is shown; 100% is the amount of label found at 180 min in the monolayers not receiving the unlabeled protein. Panels
C and D, apical surface (panel C) and
intracellular LPL (panel D). The amounts of
125I-LPL on the apical endothelial cell surface of the
experiments described in panels A and B are
shown. After removing the medium from both the upper and lower
chambers, the cells were washed with phosphate-buffered saline and then
incubated for 30 min at 4 °C with 100 units/ml heparin included in
DMEM-BSA on the apical side of the cells. Thereafter, the cells were
washed, and the radioactivity remaining within the cells was determined
by NaOH treatment of the cell-containing filters. Experiments in which
an excess of unlabeled LPL was included are denoted 300 to
signify the addition of 300 µg/ml unlabeled LPL.
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Fig. 2.
Role of LPL-HSPG interaction on LPL
transport. Effects of adding heparin on
125I-LPL transport to the upper chamber. 1 µg/ml
125I-LPL was added to the lower chamber in the presence of
heparin, and radioactivity appearing in the upper chamber was assessed.
Both 5 and 50 units/ml heparin decreased the amount of LPL.
Inset, the basolateral side of BAECs was treated with
heparinase/heparinitase (5 units each) prior to the addition of
125I-LPL. Shown are the amounts of radioactive LPL
appearing in the medium on the apical side of the monolayers after
3 h. The results are the averages ± S.D. of experiments
performed in triplicate.
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Fig. 3.
Effects of RAP on LPL transport across
endothelial cell monolayers. Panel A, effects of
different concentrations of RAP. The basolateral side of the cultured
monolayers was incubated with different concentrations of RAP (0.33-10
µg/ml) for 1 h at 37 °C. 1 µg/ml 125I-LPL was
then added to the medium in the lower chamber. The cells were incubated
at 37 °C, and 125I-LPL transported to the upper chamber
was determined. Data are the means ± S.D. of experiments
performed in triplicate; the control is without RAP. Panel
B, the addition of heparin and unlabeled LPL to RAP treated cells.
5 µg/ml RAP was added to cells, and 125I-LPL transport in
the presence of 5 units/ml heparin or an excess of unlabeled LPL (300 µg/ml) was assessed. In this experiment, as in panel A,
RAP decreased transport ~50%; neither heparin nor unlabeled LPL
greatly increased the inhibition of 125I-LPL transport.
Panel C, [3H]dextran movement across
monolayers. 1 µg/ml dextran was included in the lower chamber, and
the amount of dextran in the upper chamber was monitored over time.
Similar studies were performed using monolayers treated with RAP or
heparin as described in panel B. Panel D,
movement of 125I-LPL from the apical to basolateral side of
endothelial monolayers. 1 µg of 125I-LPL was added to the
upper chamber, and the transfer of radioactivity to the lower chamber
was monitored over time. Shown is the amount of 125I-LPL
transported across BAECs under control conditions and using cells in
which the apical surface was treated with 5 µg/ml RAP or
heparinase/heparinitase.
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Fig. 4.
Effects of heparin and RAP on cell surface
and intracellular 125I-LPL. The amounts of LPL in the
cells and on the surface of monolayers treated as described in Figs.
1-3 were determined. After a 3-h incubation, media were removed, cells
washed, and cell surface LPL was released by treating the apical
surface with DMEM-BSA containing 100 units/ml heparin at 4 °C for 30 min. After heparin treatment of the basolateral side of the cells, the
cells were washed and then dissolved in 0.1 N NaOH.
Radioactivity in the NaOH fraction represents the intracellular LPL.
Open bar, control; filled bar, treated with 5 µg/ml RAP; lined bar, treated with 5 units/ml heparin; and
dotted bar, RAP + heparin. Values represent the average ± S.D. of experiment performed in triplicate.
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Fig. 5.
Transport of LPL activity across endothelial
monolayers. 100 µg/ml purified bovine LPL was added to the lower
chamber alone or with 5 units/ml heparin (Heparin), 5 µg/ml RAP (LPL+RAP), or after incubation of the monolayers
with 5 µg/ml RAP in the bottom chamber (RAP treated). This
figure shows the lipolytic activity of 30 µl of the medium from the
upper chambers after a 1-h incubation at 37 °C. Activity was
measured as described under "Experimental Procedures." Values
represent the means of three wells for each condition ± S.D. The
lipolytic activity of each well was determined by assays run in
triplicate.
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Fig. 6.
Transcytosis of heat-inactivated
125I-LPL. Control LPL (denoted Active LPL)
or 1 µg/ml heat-inactivated 125I-LPL (denoted
Inactive LPL) was added to the basolateral side of an
endothelial cell monolayer, and the amount of 125I-LPL in
the upper chamber was determined after a 3-h incubation. In some
chambers the endothelial cell monolayers were first treated with 5 µg/ml RAP for 1 h; in other chambers the inactive LPL was
incubated with RAP prior to its inclusion in the lower chamber (denoted
Inactive LPL+RAP). Data are the means ± S.D. of
experiments performed in triplicate; the amount of nonheated
125I-LPL found in the upper chamber is shown as
100%.
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Fig. 7.
Effects of anti-receptor antibodies on LPL
transport. Panel A, antibody effects on
125I-LPL transport. Endothelial cell monolayers were
incubated with different antibodies (30 µg/ml) added to the medium on
the basolateral side for 1 h at 37 °C after which 1 µg/ml
125I-LPL was added, and 125I-LPL movement to
the upper chamber was determined. Panel B, antibody effects
on LPL activity transport. Endothelial cell monolayers were
incubated ± 30 µg/ml anti-VLDLr antibody, 5 µg/ml RAP, or
anti-VLDLr antibody and RAP together. After 1 h, 100 µg/ml
purified bovine LPL was added to the lower chamber, and the cells were
incubated at 37 °C for 1 h. LPL activity in the upper chamber
was determined. Values represent the average ± S.D. of two
experiments performed in triplicate.
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Fig. 8.
VLDLr overexpression in endothelial cell
monolayers. Panel A, RAP-binding proteins in control
and virus-infected BAECs. Cell extracts prepared from control and
virus-infected BAECs were used for SDS-polyacrylamide gel
electrophoresis on 5% gels under nonreducing conditions and
transferred to a nitrocellulose membrane. RAP-binding proteins were
visualized as described under "Experimental Procedures." Lane
1 shows control BAECs; lane 2 shows cells infected with
AdLacZ (LacZ); lane 3 shows cells infected with
AdhVLDLr (VLDLr). Panel B, transport of
125I-LPL across control and virus-infected BAECs. 1 µg/ml
125I-LPL was added to DMEM and 1.5% BSA on the basolateral
medium of control or infected BAECs, and the amount transported to the
upper chamber medium after a 3-h incubation at 37 °C is shown.
AdhVLDLr (denoted VLDLr), but not AdLacZ (denoted
LacZ), infected cells had more LPL transcytosis, and this
increase was completely inhibited by RAP. Data are the means ± S.D. of experiments performed in triplicate. Panels C and
D, effects of VLDLr expression on apical cell surface
(panel C) and intracellular (panel D) LPL. The
amounts of intracellular and apical surface LPL recovered from control
and infected cells ± RAP are shown. Data are the means ± S.D. of experiments performed in triplicate.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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ACKNOWLEDGEMENT |
---|
We are indebted to Dr. Joachim Herz who suggested a number of the preliminary experiments from which these studies originated and who shared unpublished data on the in vivo effects of RAP on LPL and lipoprotein metabolism.
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FOOTNOTES |
---|
* This work was supported in part by Grants HL-45095 (to I. J. G.), HL-03323 (to J. C. O.), and HL-50784 and HL-54710 (to D. K. S.) from the NHLBI, National Institutes of Health.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.
§ Recipient of Research Grant Ra 914/1-1 from the Deutsche Forschungsgemeinschaft.
** To whom correspondence should be addressed: Dept. of Medicine, Columbia University, 630 West 168 St., New York, NY 10032. Tel.: 212-305-3678; Fax: 212-305-5384; E-mail: ijg3@columbia.edu.
Published, JBC Papers in Press, December 19, 2000, DOI 10.1074/jbc.M008813200
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ABBREVIATIONS |
---|
The abbreviations used are:
LPL, lipoprotein
lipase;
LDL, low density lipoprotein;
LRP, LDL receptor-related
protein;
VLDL, very low density lipoprotein;
VLDLr, VLDL receptor;
HSPG, heparan sulfate proteoglycan;
RAP, receptor-associated protein;
BAEC, bovine aortic endothelial cell;
DMEM, Dulbecco's modified
Eagle's medium;
BSA, bovine serum albumin;
AdhVLDLr, human VLDLr
expressing adenovirus;
AdLacZ, -galactosidase expressing adenovirus;
IL-8, interleukin-8.
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REFERENCES |
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