The complex of the insect LDL receptor homolog, lipophorin receptor, LpR, and its lipoprotein ligand does not dissociate under endosomal conditions

Authors


K. W. Rodenburg, Division of Endocrinology and Metabolism, Department of Biology and Institute of Biomembranes, Utrecht University, NL-CH Utrecht, the Netherlands
Fax: +31 30 253 2837
Tel: +31 30 253 9331
E-mail: c.w.rodenburg@uu.nl

Abstract

The insect low-density lipoprotein (LDL) receptor (LDLR) homolog, lipophorin receptor (LpR), mediates endocytic uptake of the single insect lipoprotein, high-density lipophorin (HDLp), which is structurally related to LDL. However, in contrast to the fate of LDL, which is endocytosed by LDLR, we previously demonstrated that after endocytosis, HDLp is sorted to the endocytic recycling compartment and recycled for resecretion in a transferrin-like manner. This means that the integrity of the complex between HDLp and LpR is retained under endosomal conditions. Therefore, in this study, the ligand-binding and ligand-dissociation capacities of LpR were investigated by employing a new flow cytometric assay, using LDLR as a control. At pH 5.4, the LpR–HDLp complex remained stable, whereas that of LDLR and LDL dissociated. Hybrid HDLp-binding receptors, containing either the β-propeller or both the β-propeller and the hinge region of LDLR, appeared to be unable to release ligand at endosomal pH, revealing that the stability of the complex is imparted by the ligand-binding domain of LpR. The LpR–HDLp complex additionally appeared to be EDTA-resistant, excluding a low Ca2+ concentration in the endosome as an alternative trigger for complex dissociation. From binding of HDLp to the above hybrid receptors, it was inferred that the stability upon EDTA treatment is confined to LDLR type A (LA) ligand-binding repeats 1–7. Additional (competition) binding experiments indicated that the binding site of LpR for HDLp most likely involves LA-2–7. It is therefore proposed that the remarkable stability of the LpR–HDLp complex is attributable to this binding site. Together, these data indicate that LpR and HDLp travel in complex to the endocytic recycling compartment, which constitutes a key determinant for ligand recycling by LpR.

Abbreviations
apoLp-I

apolipophorin I

apoLp-II

apolipophorin II

CHO

Chinese hamster ovary

EGF

epidermal growth factor

ERC

endocytic recycling compartment

FACS

fluorescence-activated cell sorter

FITC

fluorescein isothiocyanate

HDLp

high-density lipophorin

LA

low-density lipoprotein receptor type A

LDL

low-density lipoprotein

LDLR

low-density lipoprotein receptor

LpR

lipophorin receptor

LRP

low-density lipoprotein receptor-related protein

OG

Oregon green

PCSK9

proprotein convertase subtilisin type 9

R1

region 1

RAP

receptor-associated protein

Lipoproteins are used to transport lipids in the circulation of vertebrates as well as invertebrates. Whereas mammals employ an array of different lipoproteins, insects rely on one single multifunctional lipoprotein, high-density lipophorin (HDLp), which is synthesized in the fat body and released into the blood (hemolymph). In addition to lipid, the particle harbors two nonexchangeable apolipoproteins, apolipophorin I (apoLp-I) and apolipophorin II (apoLp-II), which are derived from the post-translational cleavage of their common precursor, apoLp-II/I [1,2]. ApoLp-II/I was demonstrated to be a homolog of apolipoprotein B-100 [3,4], the nonexchangeable apolipoprotein of mammalian lipoproteins such as very-low-density lipoprotein and low-density lipoprotein (LDL) [5,6]. Despite this homology, HDLp appears to function differentially from these mammalian lipoproteins, as upon conversion of very-low-density lipoprotein to LDL, the latter is endocytosed by the LDL receptor (LDLR) and subsequently lysosomally degraded [7], whereas the insect lipoprotein iteratively loads and unloads lipid at various target tissues without being internalized or degraded, and thus acts as a reusable shuttle [8–11]. However, in apparent contrast to this concept of HDLp as a reusable shuttle, receptor-mediated endocytic uptake of HDLp was discovered in fat body tissue of larval and young adult locusts [12], and was shown to be mediated by an LDLR homolog [13]. This first lipophorin receptor (LpR) was molecularly and functionally characterized [13–18], and since then the LpR sequences of several other insect species have been reported [19–24]. Sequence analysis showed that LpR is a classic LDLR family member, encompassing all the typical domains in an LDLR-like sequential manner [13]: (a) a ligand-binding domain consisting of LDLR type A (LA) repeats; (b) an epidermal growth factor (EGF) precursor homology domain composed of two EGF repeats (EGF-A and EGF-B), a β-propeller containing YWTD repeats, and a third EGF repeat (EGF-C); (c) an O-linked glycosylation domain; (d) a transmembrane domain; and (e) an intracellular C-terminal domain [25]. Three-dimensional models of the elements representing the ligand-binding domain and EGF precursor homology domain of locust LpR bear a striking resemblance to those of mammalian LDLR [10]. On the other hand, the ligand-binding domain of LpR contains one additional LA repeat as compared to the cluster of seven repeats in LDLR [13,14], and despite their pronounced structural similarity, the specificity of LpR and LDLR for their ligands (HDLp and LDL, respectively) is mutually exclusive [14].

Remarkably, however, when the functioning of LpR was compared directly with that of LDLR in a mammalian cell line [Chinese hamster ovary (CHO) cells transfected with LpR], the insect lipoprotein, in contrast to LDL, was shown to remain colocalized with its receptor and was targeted to the endocytic recycling compartment (ERC). From the latter compartment, HDLp is resecreted, following a recycling pathway similar to that of transferrin [14]. In the insect system, LpR appeared to function similarly. Although an insect fat body cell line is not available, HDLp internalized by fat body tissue from young adult locusts endogenously expressing LpR is also resecreted, consistent with the above concept of ligand recycling in LpR-transfected CHO cells [12,18]. Trafficking of ligand to the ERC constitutes a highly unusual property among LDLR family members, and in addition, LDLR mutants that remain in complex with LDL are targeted to lysosomes [17,26].

During LDLR-mediated endocytosis of LDL, the receptor–ligand complex ends up in early endosomes that have a lumenal pH of 6–6.5 [27]. At this acidic pH, the ectodomain of LDLR, composed of the ligand-binding domain, EGF precursor homology domain and glycosylation domain, is proposed to undergo a conformational change, resulting in the release of bound LDL. In this model, the ligand-binding domain is hypothesized to fold onto the β-propeller after protonation of His residues located at the interface of LA-4, LA-5 and the β-propeller [28], whereas other residues at this interface, i.e. Gln540, Glu581, and Lys582, are important for docking of the ligand-binding domain onto the β-propeller [29]. In addition, the linker between LA-7 and EGF-A was demonstrated to constitute a rigid structure stabilized by a cluster of hydrophobic residues that includes Phe261, Val274, and Ile313 [29]. Because of the rigidity of this linker as well as that between EGF-A and EGF-B, the three repeats serve as a rigid scaffold, providing a favorable overall topology that permits the ligand-binding domain to fold over the β-propeller [29,30]. As a result of this conformational change, the β-propeller displaces bound LDL [28,31]. In addition to the low pH, there is a drop in Ca2+ concentration in the endosome [32], which is predicted to destabilize the Ca2+-binding properties of the LA repeats and of EGF-A and EGF-B, and thus might additionally contribute to the pH-dependent ligand release [25,33–35]. In the LA repeats, which consist of approximately 40 amino acids and are organized in a two-loop conformation stabilized by three disulfide bonds, a Ca2+ is chelated by a conserved stretch of acidic amino acids (DCxDxSDE) and is essential to stabilize the C-terminal fold of the repeat [34,36–39]. Consequently, removal of Ca2+ abolishes ligand binding by LDLR [40]. Whereas the released LDL is targeted to lysosomes for degradation [31], LDLR is directed to the ERC, and from there is efficiently recycled to the plasma membrane for another round of endocytosis [7,41].

As, in contrast to the different fates of LDLR and LDL, LpR and HDLp are both directed to the ERC, functional studies with LpR–LDLR hybrid receptors were performed to determine the molecular mechanism of LpR-mediated ligand sorting and subsequent recycling. The data obtained indicate that the ability of LpR to deliver HDLp to the ERC is not attributable to the C-terminal intracellular domain, both the length and sequence of which are very different from that of LDLR, but appears to be a function of the ectodomain [16]. The mechanism of HDLp recycling by LpR implies that during its intracellular itinerary, the LpR–HDLp complex is not dissociated. Therefore, in this study, a novel binding assay using flow cytometry was used to demonstrate that, in contrast to what is found with control experiments involving LDLR and LDL, LpR and HDLp remain in complex at endosomal pH. This remarkable stability of the receptor–ligand complex appeared to be accounted for by the ligand-binding domain. In addition, treatment of the LpR–HDLp complex with an EDTA-containing buffer to mimic the effect of the low Ca2+ concentration in the endosome did not induce complex dissociation either, once again in contrast to the LDLR–LDL complex. Together, our new findings provide ample evidence that endosomal conditions fail to result in dissociation of the complex, signifying that HDLp and LpR travel in complex to the ERC. Experiments using an LpR–LDLR hybrid receptor containing LA-1–7 of LpR and the complementary part of LDLR suggest that the stability of the complex is imparted by LA-1–7, which were shown to comprise the binding site for HDLp. The data accumulated imply that the stability of the complex is engendered by the specific interaction between LpR and HDLp.

Results

Measurement of ligand binding and subsequent endocytosis by flow cytometry

To assess the binding of HDLp to LpR, a flow cytometric assay was developed in which living, attached LDLR-deficient CHO cells [42] transfected with LpR were incubated with Oregon green (OG)-labeled HDLp (OG–HDLp). The analysis of bound ligand was performed using flow cytometry, which requires cells in suspension. Therefore, a three-step procedure was used. The first step involved the binding of ligand at 4 °C to prevent endocytosis, allowing the binding to reach equilibrium. Second, the cells were incubated at 37 °C for 5 min in serum-free medium to mediate endocytosis of bound ligand by LpR [14], protecting bound ligand from the trypsin treatment applied in the next step. The third step involved resuspension of the cells by trypsinization and measurement of the fluorescence by flow cytometry. This measurement resulted in a dotplot displaying two populations of cells with different fluorescence intensities (Fig. 1A): first, a small population with a relatively high fluorescence intensity (Fig. 1A, population 1), representing LpR-transfected cells that bound and subsequently endocytosed HDLp; and second, a population of LpR-transfected cells with a lower fluorescence intensity (Fig. 1A, population 2). The second population is located at the same position as the negative control cells (Fig. 1B), indicating that these cells did not bind HDLp. Detection of the receptor on the plasma membrane, enabled by the use of antibody 2189/90, yielded a similar distribution of the two populations (Fig. 1C, cf. Fig. 1A). Quantification of the number of cells in population 1 revealed that the number of cells that bound ligand was 91.5 ± 6.3% of the number that bound antibody, indicating that binding of HDLp was proportional to the amount of receptor on the plasma membrane.

Figure 1.

 FACS analysis of HDLp binding by LpR. After binding and subsequent endocytosis of OG–HDLp, the cells were trypsinized and analyzed by flow cytometry (A). The amount of fluorescence is plotted on the y-axis (relative values), and the forward scatter (relative values) on the x-axis. Cells in the population indicated by 1 (population 1) are transfected cells with a higher fluorescence intensity than the cells in the population indicated by 2 (population 2). (B) A similar experiment using untransfected cells. (C) Measurements of cells that were incubated at 4 °C with antibody 2189/90, and then with an FITC-labeled secondary antibody at 4 °C, and then at 37 °C for 5 min before trypsinization and analysis. The data shown in the plots are representative of four independent experiments.

HDLp and LpR remain in complex at endosomal pH

To investigate whether the LpR–HDLp complex dissociates upon exposure to endosomal pH, OG–HDLp was bound at neutral pH (7.4) to LpR-transfected cells, after which the cells were washed at 4 °C with a buffer of low pH (5.4). After endocytosis of bound ligand, the fluorescence of the cells appeared to be not affected when compared to cells that had been washed at pH 7.4 (Fig. 2A,B). In contrast, similar experiments performed with cells transfected with LDLR cDNA, which were incubated with OG–LDL and subsequently washed at pH 5.4, resulted in a decrease in fluorescence in comparison to cells that had been washed at pH 7.4 (Fig. 2C,D). After washing at pH 5.4, the population with low fluorescence intensity was located at the same position as after washing at pH 7.4, indicating that the different pH values of the buffers used in the incubations did not affect the size or the morphology of the cells, and thereby the amount of fluorescence. The population with low fluorescence intensity was excluded from the analysis by defining region 1 (R1) (Fig. 2). To quantify the amount of bound ligand, the mean fluorescence in R1 (Fig. 2) was determined and compared to the mean fluorescence in R1 after washing at pH 5.4. In the case of LpR, the fluorescence of cells washed at pH 5.4 was 94.3 ± 7.6% of the fluorescence of cells washed at pH 7.4, indicating that OG–HDLp remained bound to LpR upon exposure to pH 5.4. As expected, in the case of LDLR, the fluorescence of cells washed at pH 5.4 had decreased significantly, and amounted to only 51.4 ± 2.2% of the fluorescence of cells washed at pH 7.4 (Fig. 2E). A longer incubation at pH 5.4 (1 h instead of 30 min; data not shown) did not result in further dissociation of either the LDLR–LDL or LpR–HDLp complexes, suggesting that an incubation time of 30 min was sufficient to achieve maximum dissociation. Furthermore, the expression level of receptor, which varied between different cell lines and thus between different experiments, did not influence the relative amount of dissociation (data not shown).

Figure 2.

 LpR and HDLp remain in complex at endosomal pH. CHO(LpR) cells were incubated with OG–HDLp and washed at pH 7.4 (A) or pH 5.4 (B). (C, D) Similar experiment for binding of OG–LDL to cells transfected with LDLR, and washed at pH 7.4 (C) or pH 5.4 (D). Plots are representative of at least eight independent experiments performed on cell lines created by four different transfections. (E) Amount of bound ligand after washing at pH 5.4. The mean fluorescence (y-mean) in R1 (A–D) was determined for each sample. The relative y-mean (Rel. y-mean) after washing at pH 5.4 was calculated by the formula y-meanpH 5.4/y-meanpH 7.4, and is plotted on the y-axis. Data are the means of at least eight independent experiments. Error bars indicate the SEM. See legend to Fig. 1 for more details.

Exposure of the LpR–HDLp complex to pH values between 4.0 and 5.0 resulted in a substantial decrease in fluorescence of the cells (data not shown). However, at this pH, HDLp appeared to be precipitated (data not shown). Moreover, because in LpR-transfected CHO cells HDLp is transported from the early endosome to the ERC, and from there is returned to the plasma membrane [14], it is unlikely that the LpR–HDLp complex encounters a pH lower than 5.4. Additionally, it should be noted that the pH of endosomes in the insect fat body is similar to that of mammalian cells [43], indicating that also after LpR-mediated uptake of HDLp in insect fat body tissue the LpR–HDLp complex does not encounter a pH lower than 5.4.

The lack of LpR–HDLp complex dissociation is caused by the ligand-binding domain

Sequence alignment of the amino acid sequence of LDLR with that of LpR revealed that several of the residues crucial for LDL release by LDLR are not conserved in LpR (Table 1). To investigate whether the deficiency of these crucial residues in LpR may be responsible for the lack of dissociation of the LpR–HDLp complex, the binding and dissociation capacities of different hybrid receptors (Fig. 3A [16]) were assessed. LDLR(1–292)LpR(343–850) was able to bind LDL, but unable to release this ligand at endosomal pH (Fig. 3B), suggesting that the absence of Gln540, His562, Glu581 and Lys582 in the β-propeller of LpR causes the lack of HDLp release by LpR. However, the reciprocal hybrid, LpR(1–342)LDLR(293–839) (Fig. 3A), appeared to be equally incapable of releasing its ligand, HDLp (Fig. 3B). The presence of Gln540, His562, Glu581 and Lys582 in this hybrid suggests that the β-propeller of this hybrid may able to interact with the ligand-binding domain. However, although LpR(1–342)LDLR(293–839) contains Ile313, it does not contain the complete hinge region of LDLR. The presence of two Gly residues in LpR at the corresponding positions of His264 and Ser265 of LDLR (Table 1) might decrease the rigidity of the hinge region of LpR(1–342)LDLR(293–839), thereby abolishing ligand release. To investigate whether the complete hinge region and β-propeller of LDLR were able to induce HDLp release by LpR, the hybrid receptor LpR(1–301)LDLR(252–839) (Fig. 3A) was created. This hybrid receptor was able to bind HDLp, but, like wild-type LpR, was unable to release it (Fig. 3B). As these functional LDLR domains failed to evoke HDLp release, the lack of dissociation of the complex is proposed to result from the specific binding interaction of the ligand-binding domain of LpR with HDLp.

Table 1.   LDLR amino acid residues that are essential for LDL release and the corresponding amino acid of LpR.
Residue in LDLRCorresponding residue in LpRLocationFunctionRef.
His190His270LA-5Interaction with β-propeller[29,44]
Phe261Phe344LA-7Required for rigidity of the hinge region[29]
His264Gly346LA-7Unknownhttp://www.ucl.ac.uk/fh/
Ser265Gly347LA-7Unknownhttp://www.ucl.ac.uk/fh/
Val274Val357LA-7Required for rigidity of the hinge region[29]
Ile313Ala395EGF-AAnchorage of EGF-A and EGF-B with LA-7[29,49]
Gln540Lys621β-PropellerInteraction with ligand-binding domain[29,49]
His562Asn643β-PropellerInduction of conformational change[29]
Glu581Pro662β-PropellerInteraction with ligand-binding domain[29,49]
Lys582Glu663β-PropellerInteraction with ligand-binding domain[29]
His586His667β-PropellerInteraction with ligand-binding domain[29,44]
Figure 3.

 Hybrid receptors and relative amount of pH-dependent ligand dissociation. (A) Schematic models of the hybrid receptors. LDLR domains are depicted in gray and LpR domains in black. Each receptor contains a ligand-binding domain composed of LA repeats (squares), an EGF-precursor homology domain composed of two EGF repeats (diamonds) that are separated from a third by a β-propeller containing YWTD repeats (circle), an O-linked glycosylation domain (oval), a transmembrane domain (trapezoid), and an intracellular C-terminal domain (long rectangle). The wide and open rectangle represents the plasma membrane. The numbers indicate the amino acids of the mature proteins. Amino acids that are important for LDL release and not conserved in LpR are indicated by white dots. (B) Amount of ligand bound to different hybrid receptors after incubation at pH 5.4. CHO cells transfected with the different hybrid receptors were incubated with OG–LDL [LDLR and LDLR(1–292)LpR(343–850)] or OG–HDLp [LpR, LpR(1–342)LDLR(293–839), LpR(1–301)LDLR(252–839)] and washed at pH 7.4 or pH 5.4. The fluorescence was measured by flow cytometry. The y-mean of the receptor-expressing population was determined for each sample. The relative y-mean (Rel. y-mean) after a wash at pH 5.4 was calculated by the formula y-meanpH 5.4/y-meanpH 7.4, and is plotted on the y-axis. The values represented are the averages of at least three independent experiments. Error bars indicate the SEM. See legend to Fig. 1 for more details.

HDLp binding to LpR is not sensitive to EDTA

Ligand binding by LDLR family members is known to be dependent on Ca2+ [33,44], and the removal of Ca2+ from LDLR, e.g. by EDTA, prevents ligand binding [40]. To investigate whether the drop in Ca2+ level that occurs in the early endosome could result in a disruption of the interaction between LpR and HDLp, LpR-transfected cells that had bound OG–HDLp were exposed to an EDTA-containing buffer (Fig. 4A,B). After washing and subsequent endocytosis of bound ligand, the fluorescence of the cells was measured by flow cytometry. The mean fluorescence of cells that bound OG–HDLp did not change upon EDTA treatment, as 96.6 ± 7.5% of OG–HDLp remained bound to the receptor (Fig. 4E), demonstrating that the complex was not disrupted. In contrast, when the same experimental approach was employed using the LDLR–LDL complex as a control, this resulted, as expected, in a significant decrease of receptor-bound LDL fluorescence (Fig. 4C,D). Only 37.8 ± 4.1% of OG–LDL remained bound to the receptor (Fig. 4E). This indicates that the low Ca2+ concentration in the early endosome is not able to induce dissociation of the LpR–HDLp complex. To determine whether the stability of the complex upon EDTA treatment is caused by the LA repeats or by the two N-terminal EGF repeats (EGF-A and EGF-B), which also contain a Ca2+, similar experiments were performed with the hybrid receptors (Fig. 3A). The interaction between HDLp and LpR(1–342)LDLR(293–839) was not abrogated by EDTA treatment (Fig. 4E). As this receptor contains EGF-A and EGF-B of LDLR, this suggests that the stability results from the ligand-binding domain of LpR. In addition, the binding of HDLp to LpR(1–301)LDLR(252–839) was shown to be EDTA-resistant, indicating that the resistance resides in the first seven LA repeats of LpR. In contrast, the binding of LDL to the reciprocal hybrid receptor LDLR(1–251)LpR(302–850) (Fig. 4F), the ligand-binding domain of which is composed of the six most N-terminal LA repeats of LDLR and LA-8 of LpR, was not EDTA-resistant (Fig. 4E). Collectively, these results indicate that the EDTA resistance of the binding of HDLp to LpR is imparted by LA-1–7.

Figure 4.

 HDLp remains in complex with LA-1–7 of LpR after EDTA treatment. CHO(LpR) cells were incubated with OG–HDLp and washed at pH 7.4 without (A) or with (B) EDTA. (C, D) A similar experiment for binding of OG–LDL to cells transfected with LDLR, washed in the absence (C) and presence (D) of EDTA. (E) Amount of OG–HDLp bound to LpR, LpR(1–342)LDLR(293–839) or LpR(1–301)LDLR(252–839) or of OG–LDL bound to LDLR (control) and LDLR(1–251)LpR(302–850) after washing with EDTA. The mean fluorescence (y-mean) in R1 was determined for each sample. The relative y-mean (Rel. y-mean) after a wash with an EDTA-containing buffer was calculated by the formula y-meanEDTA/y-meanpH 7.4, and is plotted on the y-axis. Data are the means of at least six independent experiments. Error bars indicate the SEM. See legend to Fig. 1 for more details. (F) Schematic model of LDLR(1–251)LpR(302–850). LDLR domains are depicted in gray and LpR domains in black. See legend to Fig. 3 for more details.

HDLp binding by LpR is similar to ligand binding by other LDLR family members

As neither treatment with EDTA nor endosomal pH was able to disrupt the complex, the issue was addressed of whether LpR binds HDLp in a different manner than other LDLR family members bind their ligands. Therefore, the ability of receptor-associated protein (RAP), a general ligand for LDLR family members [45–47], to compete with HDLp binding to LpR was assayed. LpR-transfected cells incubated with OG–HDLp in the presence of an equimolar concentration of RAP displayed a fluorescence similar to that of untransfected cells incubated with OG–HDLp and RAP (Fig. 5), indicating that RAP completely blocked the binding of OG–HDLp to LpR. Thus, RAP and HDLp apparently use the same binding site. Therefore, LpR probably binds HDLp using the general mechanism of binding of ligands by LDLR family members [48,49].

Figure 5.

 RAP competes with OG–HDLp for binding. CHO(LpR) cells were incubated with OG–HDLp (A) or OG–HDLp in the presence of RAP (B). (C, D) A similar experiment using untransfected cells incubated with OG–HDLp in the absence (C) or presence (D) of RAP. Plots are representative of three independent experiments. See legend to Fig. 1 for more details.

LA-8 and EGF-A of LpR are not involved in the binding site of LpR for HDLp

To characterize the binding site for HDLp in LpR, the binding of ligand by wild-type LpR was compared with the binding of HDLp by LpR(1–301)LDLR(252–839). To exclude the possibility that differences in ligand binding were caused by differences in receptor expression, binding of antibody to the receptor was used as a measure for the amount of receptor on the plasma membrane. After binding, cells were allowed to endocytose bound ligand or antibody, by incubation of the cells at 37 °C. Following endocytosis, the cells were trypsinized, and the fluorescence was analyzed with flow cytometry. As a control, the binding of LDL to LDLR was compared to the binding of LDL to LDLR(1–251)LpR(302–850). For wild-type LDLR, both ligand binding and antibody binding yielded similar numbers of fluorescent cells in R1 (Table 2), indicating that the amount of LDL binding was proportional to the amount of LDLR on the plasma membrane. However, in the case of the hybrid receptor LDLR(1–251)LpR(302–850) (Fig. 4F), the ligand-binding domain of which is composed of the six most N-terminal LA repeats of LDLR and LA-8 of LpR, the number of cells that bound ligand was only 58.7 ± 4.2% of the number of cells that bound antibody. This suggests a reduction in affinity of the hybrid receptor for LDL, which may be expected, as the binding site of LDLR for LDL encompasses LA-3–7 and EGF-A [50,51]. Despite the presence of LA-8 of LpR in this receptor, LDLR(1–251)LpR(302–850) was not able to bind HDLp. Moreover, as was previously found for ligand binding to LpR and LDLR [14], for binding to the hybrid receptors the ligands are not interchangeable (data not shown) [16]. With respect to the binding of HDLp to LpR, the number of cells that bound ligand was 91.5 ± 6.3% of the number of cells that bound antibody, showing that also for LpR the binding of HDLp is proportional to LpR expression. As for the hybrid receptor LpR(1–301)LDLR(252–839), which contains LA-1–7 of LpR, followed by LA-7 of LDLR, the binding of HDLp yielded 84.4 ± 7.5% of fluorescent cells as compared to the number of cells that bound antibody (Table 2). As 84.4 ± 7.5% is not significantly lower than the 91.5 ± 6.3% measured for LpR, this suggests that LpR(1–301)LDLR(252–839) binds HDLp with a similar affinity as wild-type LpR. These results indicate that LA-7 and EGF-A of LDLR were able to replace the corresponding region of LpR (LA-8 and EGF-A) without disrupting the binding site for HDLp. This suggests that these repeats of LpR are not involved in the ligand-binding site of LpR, in contrast to the same structure (LA-7 and EGF-A) in LDLR.

Table 2.   Binding efficiency of expressed receptors. CHO cells transfected with the different (hybrid) receptors were incubated with OG–LDL [LDLR, LDLR(1–251)LpR(302–850)] or with OG–HDLp [LpR, LpR(1–301)LDLR(252–839), LpRsplice] or a primary antibody detected by an FITC-labeled secondary antibody. The percentage of cells that bound ligand relative to the percentage of cells that bound antibody was determined. The percentages shown are the means ± SEM of at least five independent experiments.
ReceptorBinding efficiency (%)
LDLR96.1 ± 7.0
LDLR(1–251)LpR(302–850)58.7 ± 4.2
LpR91.5 ± 6.3
LpR(1–301)LDLR(252–839)84.4 ± 7.5
LpRsplice57.3 ± 5.9

LA-3 is involved in the binding site of LpR for HDLp

Recently, a putative splice variant of LpR, LpRsplice, has been identified in ovaries of young animals (J. Kerver and K. W. Rodenburg, unpublished results), in which the sequence of LA-3 is altered. Although sequence alignment revealed a high similarity between LA-3 of the two variants, the central Trp present in LA-3 of wild-type LpR is absent in LA-3 of LpRsplice (Fig. 6). As the central Trp plays an important role in the interaction between LDLR family members and their ligands [49], we investigated whether the binding of OG–HDLp to LpRsplice deviates from that to wild-type LpR. The binding of HDLp to LpRsplice yielded only 57.3 ± 6.9% of fluorescent cells as compared to the number of cells that bound antibody (Table 2), implying that LpRsplice binds HDLp with a lower affinity than wild-type LpR. This indicates that LA-3 is involved in the binding of HDLp to wild-type LpR, suggesting that the Trp in wild-type LpR may be involved in the interface between HDLp and LpR.

Figure 6.

 Sequence of LA-3 of a putative splice variant in LpR. Alignment of the sequences of LA-3 of wild-type LpR and LpRsplice. Identical residues are boxed in black, and conserved residues are shaded in gray. The arrow indicates the position of the central Trp in the sequence of wild-type LpR.

LA-1 is not essential for binding of ligand to LpR

To further investigate which LA repeats are involved in HDLp binding, we investigated whether antibody 2189/90, directed against the first LA repeat of LpR, was able to compete with HDLp for binding by LpR. After a preincubation with OG–HDLp at 4 °C, followed by incubation with antibody 2189/90 at 4 °C, the fluorescence after uptake of the bound OG–HDLp appeared to be 73.2 ± 6.3% of the fluorescence of the cells in such an experiment without incubation with antibody 2189/90. In a similar experiment in which the order of the incubations with OG–HDLp and antibody 2189/90 was reversed, the fluorescence of the cells was 78.9 ± 7.0% of the fluorescence of cells incubated with OG–HDLp alone. Although the presence of antibody resulted in a significant decrease in fluorescence of the cells as compared to cells that were incubated with OG–HDLp only, these results indicate that LpR is still able to efficiently bind a major amount of OG–HDLp in the presence of the antibody. Moreover, the amount of competition was similar to that in the corresponding control experiment using LDLR and antibody C7, an antibody against the first repeat of LDLR (data not shown) [52,53]. As LA-1 of LDLR is not involved in LDL binding [51], the inhibition of binding that was measured probably results from steric hindrance due to the size of the antibody and not from competition for the same binding site. No competition was observed between RAP and the antibody (data not shown), suggesting that LA-1 is not involved in the binding of LpR to RAP or HDLp.

Discussion

Previous studies have demonstrated that in a mammalian model (CHO) cell line transfected with insect LpR, the receptor recycles its ligand, HDLp, in a transferrin-like manner, in contrast to endogenously expressed LDLR, the ligand of which (LDL) is released and undergoes intracellular degradation [14]. Also during insect development, LpR appeared to function similarly, since HDLp internalized by fat body tissue from young adult locusts endogenously expressing LpR appeared to be resecreted, supporting the concept of LpR-mediated ligand recycling [12,18]. To investigate the mechanism underlaying the highly unusual behavior of this insect LDLR family member, we examined the stability of the binding of HDLp to LpR in direct comparison with that of LDL to LDLR, and additionally explored the subset of structural features in LpR that may allow for the occurrence of the difference in ligand delivery as compared to that in mammals.

Our present studies provide the new findings that the complex of LpR and HDLp is stable at endosomal pH and EDTA-resistant, both in contrast to the complex of LDLR and LDL. This stability of the LpR–HDLp complex is proposed to be caused by the specific interaction between HDLp and LA-2–7. Together, our data indicate that the complex of LpR and HDLp remains intact during its intracellular itinerary, which is in complete agreement with the occurrence of ligand recycling [14,16–18], and may provide a vital determinant of the ligand-recycling capacity of LpR. In several studies, flow cytometry has been used to quantify lipoprotein binding and uptake [29,53–59]. In most cases, the experiments were performed on blood cells. As these cells are already in suspension, they can be easily measured by flow cytometry. In the case of attached cells, resuspending the cells may destroy the interaction between receptor and ligand or antibody. For this reason, the actual binding experiment was performed at 4 °C to prevent endocytosis, so that equilibrium binding was achieved. After binding, the cells were allowed to endocytose bound ligand to protect it from the subsequent trypsin treatment. Fluorescence images of the cells after binding at 4 °C and after endocytosis at 37 °C showed that the bound ligand or antibodies were efficiently endocytosed, indicating that the amount of intracellular fluorescence was proportional to the amount of bound ligand at equilibrium (data not shown). For these experiments, stably transfected polyclonal cell lines were used to provide heterogeneous samples of cells that express the receptor. This resulted in flow cytometry plots containing two populations, one of which comprised cells whose fluorescence did not exceed that of untransfected cells (Fig. 1). In order to analyze only the cells that expressed the receptor, R1 was defined to exclude the population with lower fluorescence intensity from the analysis (Figs 2 and 4). However, for LDLR, the decrease in fluorescence after treatment at pH 5.4 or with EDTA resulted in a decrease of the number of cells in R1. As this reduction in sample size introduced a bias into the analysis, the number of cells in the analysis was restored by using random measurements from the population with low fluorescence intensity. After correction of the mean fluorescence, similar values for LDL release by LDLR were obtained as measured by Blacklow et al. for monoclonal cell lines [29]. The relative amount of dissociation was not affected by differences in receptor expression; we therefore conclude that the results obtained with the flow cytometric assay represent physiologically relevant receptor properties.

Our data indicate that, unlike the complex of LDLR and LDL, the complex of LpR and HDLp remains stable at a pH as low as 5.4, which is significantly lower than that encountered in endosomes (pH 6–6.5) [27]. This indicates that, despite the substantial sequence similarity between LpR and LDLR, LpR is unable to release HDLp in the early endosome. LDLR is hypothesized to release LDL at endosomal pH by undergoing a conformational change in which the β-propeller displaces LDL [31]. Blacklow et al. elegantly identified domains and residues that are important for LDL release by LDLR [29,59] (Table 1). In agreement with these results, the β-propeller of LpR, lacking the important residues Gln540, His562, Glu581, and Lys582, was incapable of inducing LDL release. Similar results were obtained for the swap of the β-propeller of LDLR with β-propeller 4 of LDLR related protein (LRP) 6, in which two Lys residues and one His are not conserved. However, when β-propeller 2 of LRP6 was introduced, containing these residues, the receptor was able to release LDL, indicating that a different β-propeller is able to substitute for the wild-type propeller of LDLR [29]. However, introducing the β-propeller of LDLR into LpR did not lead to HDLp release by the hybrid receptor LpR(1–342)LDLR(293–839), implying that other domains produce the remarkable stability of the complex. In LDLR, the interface between LA-7 and EGF-A, the hinge region, also plays an important role in LDL release, as this region functions as a rigid scaffold allowing the β-propeller to fold over the ligand-binding domain. To investigate the importance of this hinge region for the lack of HDLp release by LpR, both the hinge region and β-propeller of LDLR were introduced into LpR. The resulting hybrid receptor, LpR(1–301)LDLR(252–839), did not release HDLp, despite the fact that this hybrid contains all the domains of LDLR that are essential for LDL release. This suggests that the β-propeller of LDLR is not able to compete with HDLp for binding to the ligand-binding domain of LpR, implying that the lack of HDLp release is mainly caused by the interaction between HDLp and the ligand-binding domain of LpR, and suggests that LpR may use a different mechanism to release HDLp, in contrast to the mechanism of LDL release by LDLR, in which the β-propeller is of vital importance [28,29,60]. Interestingly, our earlier localization studies of the hybrid receptors revealed that the intracellular fate of the complex is determined by the extracellular domain as a whole [16]. In view of the mechanism of ligand recycling by LpR, this implies that for the stability of the complex, the ligand-binding domain is sufficient, but for proper targeting of LpR to the ERC, the combination of the ligand-binding domain and β-propeller of LpR is essential [16].

Ligand binding to LDLR family members is known to depend on Ca2+, due to the stabilization of the LA repeats by a central Ca2+ [33,34,36–39]. Sequence comparison of the LA repeats of LpR with those of other LDLR family members, as well as modeling and molecular dynamics studies of LA-4–6 of LpR, indicate that this also applies for the LA repeats of LpR (S. D. Roosendaal, S. Cuesta-López, J. Sancho and K. W. Rodenburg, unpublished results). In addition to a decrease in pH, the Ca2+ concentration in the early endosome drops within minutes to the low micromolar range [32], possibly contributing to ligand release by LDLR [33]. Therefore, the LpR–HDLp complex was exposed to a Ca2+-chelating agent (EDTA) to mimic the effect of low Ca2+ in the early endosome. In contrast to the binding of LDL to LDLR, the binding of HDLp to LpR appeared to be resistant to EDTA treatment. A possible explanation for this phenomenon might be that LpR binds HDLp by using a different binding mode than that used by other LDLR family members for binding of their ligands. For example, a different, Ca2+-independent binding mode is used in the interaction between the single LA repeat of Tva, the cellular receptor for subgroup A Rous sarcoma virus [61] and its ligand. However, as the ligand bound Tva with an aberrant binding mode, RAP appeared to be unable to compete with the ligand for binding to Tva [62]. Our studies show that RAP efficiently competes with HDLp for binding to LpR, indicating that HDLp binds LpR using the same binding mode as other ligands for LDLR family members, which again implies the presence of Ca2+ in LA repeats of LpR. Therefore, the resistance of the LpR–HDLp complex may be caused by a higher affinity of the LA repeats of LpR for Ca2+, or by the ability of HDLp to shield the calcium ions from EDTA. Although it is unclear what precisely causes this remarkable stability, our data emerging from the use of hybrid receptors indicate that the stability of the complex at low pH and upon EDTA treatment is caused by the interaction between HDLp and LA-1–7 of LpR. The general binding mode of LDLR family members and their ligands consists of an acidic binding pocket present in the LA repeats that entraps a Lys from the ligand. The binding is augmented by an essential aromatic residue, preferentially Trp, of the LA repeat, positioned next to the binding pocket [45,47,49,63]. To obtain more information about the recognition interface between LpR and HDLp, the LA repeats of LpR were aligned with those of LDLR. This revealed that only LA-1–6 of LpR contain a central aromatic residue, in all cases Trp (data not shown). As LA-1 appeared not to be involved in the binding site for HDLp, this suggests that only LA-2–6 are involved in the interface. LA-3 of wild-type LpR contains the central Trp, but importantly, in addition to other amino acid changes, LA-3 of LpRsplice lacks this Trp (Fig. 6). As LpRsplice binds HDLp with a lower affinity, indicating that LA-3 is involved in the interaction with HDLp, it may very well be that the absence of the Trp weakens the interaction between HDLp and LpR. In this respect, it is interesting to note that the binding of HDLp to the splice variant is also resistant to low pH and EDTA treatment (data not shown). This suggests that the Trp and the other residues that are different between LA-3 of wild-type LpR and LpRsplice are not important for determining the stability of the interaction under endosomal conditions. Additionally, from these results, it is apparent that the stability of the complex at endosomal pH and upon EDTA treatment is not merely the result of the affinity of the interaction, but may require additional contacts or a slightly alternative mode of binding of HDLp to LpR. An alternative mechanism for a stable complex at endosomal pH may be provided by the binding of proprotein convertase subtilisin type 9 (PCSK9) to LDLR. PCSK9 has been shown to be involved in the regulation of cell surface LDLR levels. After endocytosis, the LDLR–PCSK9 complex is also not dissociated at endosomal pH. Instead, the affinity of PCSK9 for LDLR is enhanced by the low pH [64,65], possibly through protonation of the abundant His residues on the surface of PCSK9 [65,66]. Even though HDLp binds to the LA repeats of the receptor and PCSK9 to EGF-A of LDLR, similar effects may play a role in the stability of the complex of HDLp and LpR. An important difference is, however, that binding of PCSK9 seems to target LDLR to lysosomes for degradation [67,68], whereas the complex of HDLp and LpR is transported to the ERC for recycling.

The acidic residues involved in Ca2+ binding of specific LA repeats are proposed to interact with the basic residues of the ligand, in particular one protruding Lys [49]. A consensus sequence containing the protruding Lys was proposed, in which the Lys is surrounded by basic and hydrophobic residues [69,70]. Such sequences are numerous in both apolipoproteins of HDLp, apoLp-I and apoLp-II. Interestingly, the three-dimensional model of their protein precursor apoLp-II/I reveals that at least one of these motifs is situated at the end of an α-helix [71], as is the case for the binding site of RAP and apolipoprotein E for LRP [49]. Furthermore, this helix is probably exposed on the surface of the HDLp particle, and is thus available for interaction with LpR [71]. Because of the multitude of putative binding sites in apoLp-I and apoLp-II, it cannot excluded that one HDLp particle binds several receptors concomitantly, as is the case for apolipoprotein E-containing lipoproteins [72,73]. In this respect, it is important to note that apolipoprotein B-100, which is a homolog of apoLp-II/I [3,4], also contains several of these consensus sequences (data not shown). However, LDL binds to LDLR with a stoichiometry of 1 : 1. Moreover, RAP is able to efficiently compete at equimolar concentrations with the binding of HDLp. Although several RAP molecules may be able to bind LpR, as RAP binds to two LA repeats [49,74], competition binding studies indicated that RAP and antibody 2189/90 against LA-1 of LpR do not compete (data not shown), suggesting that RAP does not bind LA-1. On the basis of the presence of an important acidic residue [74], sequence analysis of the LA repeats of LpR suggests that RAP may bind either LA-4–5 or LA-5–6, suggesting that the stoichiometry is one RAP molecule per LpR molecule. Therefore, it seems unlikely that LpR binds more than one HDLp molecule.

In conclusion, our results indicate that the interaction between HDLp and LA-2–7 of LpR is stable upon exposure to endosomal pH as well as EDTA treatment, implying that the integrity of the complex is maintained during intracellular trafficking of LpR and HDLp in LpR-transfected CHO cells and most likely also in insect cells. Similar to transferrin recycling, the intracellular transfer of lipid or other hydrophobic compounds from or to the HDLp particle may change its affinity for LpR, thus allowing HDLp resecretion. Indeed, binding studies using a partially delipidated HDLp particle revealed that the affinity of LpR for HDLp is modulated by the amount of lipids (S. D. Roosendaal, J. M. Van Doorn, K. M. Valentijn, D. J. Van der Horst and K. W. Rodenburg, unpublished results), suggesting that changes in lipid content may trigger HDLp resecretion. The stability of the complex and the modulation thereof may be determined by secondary contacts between HDLp and nonconserved residues of LpR. Although the function of recycling of endocytosed lipoprotein ligand during insect development remains to be defined, our study reveals the molecular mechanism underlying the stability of the LpR–HDLp complex; this is likely to provide a crucial key to the process of ligand recycling, and might additionally help to explain the ability of LDLR family members to bind a wide range of structurally unrelated ligands.

Experimental procedures

Proteins and antibodies

Insect HDLp was isolated from locust hemolymph by density gradient ultracentrifugation as described previously [14]. Human LDL was isolated from blood plasma (Bloedbank Midden Nederland, the Netherlands) as described by Redgrave et al. [75], with minor adaptations to the original protocol. The salt solutions of different densities used in the procedure contained 86.89 g·mL−1 KBr (density 1.063 g·mL−1), 18.36 g·mL−1 KBr (density 1.019 g·mL−1), and 8.68 g·mL−1 KBr (density 1.006 g·mL−1). Polyclonal rabbit antibody to LpR (antibody 2189/90) was raised against a synthetic peptide representing the unique N-terminal 20 amino acids (34–53) of LA-1 of LpR [15]. Mouse antibody to LDLR (antibody C7) was a generous gift from I. Braakman (Utrecht University, Utrecht, the Netherlands). Human His-tagged RAP (RAP–His) was a generous gift from M. Etzerodt (IMSB, Aarhus University, Århus, Denmark).

Construction of expression vectors encoding lipoprotein receptor cDNA

The cloning of the expression vectors was performed according to standard laboratory procedures and according to the protocols supplied with enzymes and kits. Site-specific mutations were generated with a QuickChange site-directed mutagenesis kit using PfuTurbo DNA polymerase (Stratagene, Amsterdam, the Netherlands), according to the manufacturer’s protocol. PCR fragments were generated using PfuTurbo DNA polymerase and synthetic oligonucleotide primers (Biolegio, Nijmegen, the Netherlands). Endonucleases were from New England BioLabs (Westburg B.V., Leusden, the Netherlands) and Fermentas (St Leon-Rot, Germany). Plasmid pcDNA3–LpR(1–297)LDLR(248–839) was made as follows. First, by mutagenesis, a unique AgeI site was introduced in pcDNA3–LpR (piLR-e [13], causing a silent mutation in the Pro301 codon (CCA→CCG; the first amino acid is that of the mature protein), using the oligonucleotides 5′-GAGAATTGCACATCACCGGTGCCAAAGTGTGACCC-3′ (forward primer) and 5′-GGGTCACACTTTGGCACCGGTGATGTGCAATTCTC-3′ (reverse primer), yielding the construct pcDNA3–LpR(AgeI). Subsequently, to replace the sequence encoding LA-8 of LpR with that encoding LA-7 of human LDLR, a 1668 bp fragment containing the 5′-flanking AgeI and 3′-flanking AccIII sites was generated by PCR from pGEM-T–LDLR(1–292)LpR(343–850) [16], using the oligonucleotides 5′-GGCCGCACCGGTGACACTCTGCGAGGGACCC-3′ (forward primer) and 5′-GCGGCCGCTTATACATAATCATTTGTCCC-3′ (reverse primer). The AgeI–AccIII fragment encoding the 5′-end LA-7 of LDLR obtained by PCR was cloned in pcDNA3–LpR(AgeI), using the enzymes AgeI and AccIII, thereby replacing the sequence encoding LA-8 of LpR to yield the mosaic receptor construct pcDNA3–LpR(1–301)LDLR(252–292)LpR(343–850). Subsequently, the 1267 bp EcoRI–KpnI fragment [16] from the mosaic receptor construct was isolated and cloned into pGEM-T–LpR(1–342)LDLR(293–839) digested with the same two endonucleases to replace the sequence encoding LA-1 through LA-8 of LpR with the sequence encoding LA-1 through LA-7 of LpR combined with LA-7 of human LDLR, thereby generating pGEM-T–LpR(1–301)LDLR(252–839). Finally, the EcoRI–NotI fragment encoding the LpR(1–301)LDLR(252–839) sequence was cloned in pcDNA3 digested with the same enzymes to yield pcDNA3–LpR(1–301)LDLR(252–839).

Plasmid pcDNA3–LDLR(1–251)LpR(302–850) was constructed similarly. First, a unique HpaI site was introduced in pcDNA3–LDLR [16], causing a silent mutation in the Asn251 codon (AAT→AAC), using the oligonucleotides 5′-GGCTGCGTTAACGTGACACTCTGCGAG-3′ (forward primer) and 5′-CTCGCATGTCAGGTTAACGCAGCC-3′ (reverse primer), yielding the construct pBS–LDLR(HpaI). Subsequently, to replace the sequence encoding LA-7 of human LDLR with that encoding LA-8 of LpR, a 1790 bp fragment, containing the unique HpaI and Bsu361 sites, was generated from pcDNA3–LpR(1–342)LDLR(293–839) by PCR, using the oligonucleotide primers 5′-CCCGGGGTTAACGTGCCAAAGTGTGACCCC-3′ (forward primer) and 5′-ATTTAAATTCACGCCAGCTCATCCTCC-3′ (reverse primer). The 473 bp HpaI–Bsu361 fragment, encoding LA-8 at the 5′-end, obtained by PCR, was then cloned in pBS–LDLR(HpaI), using the enzymes HpaI and Bsu361, replacing the sequence encoding LA-7 of LDLR with that encoding LA-8 of LpR, to yield the mosaic receptor pBS–LDLR(1–251)LpR(301–342)LDLR(293–839). To obtain the construct pcDNA3–LDLR(1–251)LpR(301–342)LDLR(293–839), the sequence encoding the mosaic receptor was cloned in pcDNA3using the XbaI restriction enzyme. Subsequently, the 225 bp EcoRI–KpnI fragment from the mosaic receptor construct was isolated and cloned into pGEM-T–LDLR(1–292)LpR(343–850) digested with the same two endonucleases to replace the sequence encoding the seven LA repeats of LDLR with that encoding LA-1–6 of LDLR followed by LA-8 of LpR, thereby generating pGEM-T–LDLR(1–251)LpR(301–850). Finally, the HindI–NotI fragment encoding the LDLR(1–251)LpR(301–850) sequence was cloned in pcDNA3 digested with the same enzymes to yield pcDNA3–LDLR(1–251)LpR(301–850). All PCR- and mutagenesis-generated LpR fragments were sequenced, and their sequences, apart from the intended mutations, were confirmed to be identical to that of LpR as indicated in the EMBL sequence database (accession number AJ000010).

Cell culture

CHO cells were cultured in 25 cm2 polystyrene culture flasks in growth medium [Ham F10 nutrient mixture (GibcoBRL, Invitrogen, Breda, the Netherlands)] containing 5% heat-inactivated fetal bovine serum (GibcoBRL) and 100 U·mL−1 penicillin G sodium and 100 μg·mL−1 streptomycin sulfate in 85% saline (GibcoBRL). The cells were maintained at 37 °C and 5% CO2.

Transfections

LDLR-deficient CHO(ldlA) cells [42] were grown up to 40% confluency in 12-well multidishes (Costar, Corning BV Life Sciences, Schiphol-Rijk, the Netherlands). After washing of the cells once, the growth medium was replaced with 500 μL of fresh growth medium. Subsequently, 2 μg of DNA and 4 μg of poly(ethylenimine) (Polysciences, Eppelheim, Germany) in 50 μL of serum-free medium (Ham F10 nutrient mixture supplemented with 100 U·mL−1 penicillin G sodium and 100 μg·mL−1 streptomycin sulfate in 85% saline) was administered to the cells. After 4 h, 500 μL of growth medium was added and cells were cultured overnight. The next day, cells were detached from dishes and cultured in 25 cm2 culture flasks in growth medium supplemented with 400 μg·mL−1 geneticin (GibcoBRL) or 400 μg·mL−1 zeocin (Cayla, Toulouse, France). Ten days after transfection, cells were used for experiments.

Fluorescence labeling of LDL and HDLp

LDL and HDLp were covalently labeled with OG 488 carboxylic acid (Molecular Probes, Leiden, the Netherlands) as described previously [14].

Binding experiments using flow cytometry

The cells were grown up to a confluency of 70%. Sixteen hours before the experiment, the growth medium was replaced with serum-free medium. At the start of the experiment, the cells were placed on ice. Subsequently, the cells were washed with ice-cold binding buffer (50 mm Tris/HCl, 2 mm CaCl2, 150 mm NaCl, pH 7.4, 4 °C) and incubated with OG-labeled LDL (35 μg·mL−1) or HDLp (25 μg·mL−1) in binding buffer for 30 min. After binding, the cells were washed once with either ice-cold binding-buffer, low-pH buffer (25 mm Tris, 25 mm sodium succinate, 2 mm CaCl2, 150 mm NaCl, pH 5.4 or pH 4.0, 4 °C) or EDTA-containing buffer (50 mm Tris/HCl, 150 mm NaCl, 5 mm EDTA, pH 7.4, 4 °C). The cells were then incubated with the buffer for 30 min. After washing of the cells, the cells were incubated for 5 min at 37 °C in serum-free medium, to allow the cells to endocytose bound ligand. After endocytosis, the cells were detached using trypsin/EDTA (Invitrogen), according to the manufacturer’s instructions, and resuspended in growth medium. Resuspended cells were fixed in 0.5% paraformaldehyde in NaCl/Pi at 4 °C for at least 30 min or overnight.

The receptor on the plasma membrane was detected using the same protocol as for ligand binding. However, the cells were incubated with an antibody against the first LA repeat (antibody C7 [76] for LDLR, and antibody 2189/90 [15] for LpR) for 30 min in binding buffer. After being washed with binding buffer, the cells were incubated for 30 min with fluorescein isothiocyanate (FITC)-labeled secondary anti-IgG (Jackson ImmunoResearch Laboratories Inc., Brunschwig, Amsterdam, the Netherlands). Then, the complex was endocytosed, and cells were detached and fixed as described above.

Competition binding experiments

Competition experiments were performed similarly to the binding experiments. However, the cells were first incubated for 30 min with primary antibody 2189/90, and then for 30 min with OG–HDLp, or vice versa. The degree of binding was compared to the degree of binding without the antibody incubation. For competition experiments with RAP, RAP–His (3.6 μg·mL−1) was added simultaneously with (OG)–HDLp or primary antibody 2189/90. Then, bound OG–HDLp or antibody 2189/90 was detected as described previously. RAP–His was detected by subsequent washing and incubation of mouse antibody to His (Amersham Biosciences, Roosendaal, the Netherlands) in binding buffer, and this was followed by washing and incubation with FITC-labeled secondary anti-IgG (Jackson ImmunoResearch Laboratories Inc.).

Flow cytometry data analysis

Samples were measured using a fluorescence-activated cell sorter (FACS; Becton Dickinson FACS Calibur). Flow cytometry data were collected using cellquest (Becton Dickinson) and downloaded into the program winmdi (TSRI FACS Core Facility, La Jolla, CA, USA) for analysis. For each sample (100 000 cells), the fluorescence was plotted against the forward scatter. On the basis of samples of untransfected cells, for each series of experiments R1 was defined to exclude cells whose fluorescence did not exceed that of untransfected cells from the analysis. Then, the number of cells and the mean fluorescence (y-mean) in R1 were determined. If the number of cells in R1 decreases with the different treatments of the cells, the y-mean in R1 is overestimated. Therefore, for each cell line, the number of cells in R1 after different treatments was compared by a t-test for paired samples performed on the logarithms of the number of cells. In cases of a significant (P < 0.05) difference in sample size due to the different treatments, the y-mean was corrected by using random values of the missing number of cells from the population with lower fluorescence intensity. After correction, for each sample the relative amount of fluorescence as compared to control samples was determined. Data presented as means ± SEM were obtained from at least three independent experiments. To test whether samples were significantly different from control samples, a t-test for paired samples was performed on the logarithms of the y-means.

Acknowledgements

We thank Steve Blacklow, Sander Meijer, Ineke Braakman, Jürgen Gent, Manon Wildenberg and Masja van Oort for stimulating discussions, Ger Arkesteijn for his help with flow cytometry, Wim Busschers for help with quantification of the flow cytometry data, and Santiago Cuesta-López and Javier Sancho for molecular dynamics studies on LA-4–6 of LpR.

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