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EMBO J. Nov 1, 2001; 20(21): 5876–5886.
PMCID: PMC125289

Identification of interaction domains of the prion protein with its 37-kDa/67-kDa laminin receptor


Cell-binding and internalization studies on neuronal and non-neuronal cells have demonstrated that the 37-kDa/67-kDa laminin receptor (LRP/LR) acts as the receptor for the cellular prion protein (PrP). Here we identify direct and heparan sulfate proteoglycan (HSPG)-dependent interaction sites mediating the binding of the cellular PrP to its receptor, which we demonstrated in vitro on recombinant proteins. Mapping analyses in the yeast two-hybrid system and cell-binding assays identified PrPLRPbd1 [amino acids (aa) 144–179] as a direct and PrPLRPbd2 (aa 53–93) as an indirect HSPG-dependent laminin receptor precursor (LRP)-binding site on PrP. The yeast two-hybrid system localized the direct PrP-binding domain on LRP between aa 161 and 179. Expression of an LRP mutant lacking the direct PrP-binding domain in wild-type and mutant HSPG-deficient Chinese hamster ovary cells by the Semliki Forest virus system demonstrates a second HSPG-dependent PrP-binding site on LRP. Considering the absence of LRP homodimerization and the direct and indirect LRP–PrP interaction sites, we propose a comprehensive model for the LRP–PrP–HSPG complex.

Keywords: heparan sulfate proteoglycans/in vitro interaction/37-kDa laminin receptor precursor/67-kDa high-affinity laminin receptor/LRP-PrP-binding domains


We recently identified the 37-kDa laminin receptor precursor (LRP) as an interactor for the prion protein (PrP) (Rieger et al., 1997; for reviews see Rieger et al., 1999; Gauczynski et al., 2001a). Employing a series of neuronal and non-neuronal cells, we proved that the 37-kDa LRP/67-kDa high-affinity laminin receptor (LR) acts as the receptor for the cellular PrP (Gauczynski et al., 2001b). In the present manuscript we used the yeast two-hybrid system and cell-binding studies on neuronal as well as non-neuronal cells involving the Semliki Forest virus (SFV) system (for reviews see Liljestrom and Garoff, 1991; Tubulekas et al., 1997) to identify domains on the PrP and the LRP involved in the PrP–LRP interaction on the cell surface. We identified two binding domains for LRP on PrP termed PrPLRPbd1 and PrPLRPbd2. The first one binds directly to LRP, whereas the second one depends on the presence of heparan sulfate proteoglycans (HSPGs) on the cell surface. The yeast two-hybrid system and cell-binding assays on wild-type and mutant HSPG-deficient Chinese hamster ovary (CHO) cells also identified two binding domains for PrP on LRP.

The relationship between 37-kDa LRP and 67-kDa LR is not yet fully understood and has been explained with homodimerization of 37-kDa LRP (Landowski et al., 1995) or an additional factor, such as a polypeptide (Castronovo et al., 1991), which might bind to 37-kDa LRP to form the 67-kDa form of the receptor. The 67-kDa heterodimer might be stabilized by hydrophobic interactions mediated by fatty acids such as palmitate, oleate and stearate bound to 37-kDa LRP and to a galectin-3 (gal-3) cross reacting polypeptide (Landowski et al., 1995; Buto et al., 1998). However, we recently proved that the β-galactoside lectin gal-3 is not present on the surface of neuronal or non-neuronal cells used for PrP-binding/internalization studies (Gauczynski et al., 2001b) and anti-gal-3 antibodies failed to compete for the 37-kDa LRP/67-kDa LR-mediated binding and internalization of the cellular PrP (Gauczynski et al., 2001b), suggesting that gal-3 is not a partner of the 37-kDa LRP in this context. In this study we investigated by a yeast two-hybrid system analysis whether gal-3 interacts with 37-kDa LRP and/or the cellular PrP. In addition, we investigated whether 37-kDa LRP interacts with itself in the yeast two-hybrid and analysed the monomer/dimer status of the receptor by size-exclusion chromatography. Both PrP (Gabizon et al., 1993; Caughey et al., 1994; Chen,S.G. et al., 1995; Brimacombe et al., 1999) and the 37-kDa/67-kDa LR (Guo et al., 1992; Kazmin et al., 2000) bind to heparan sulfates. HSPGs are required for the binding of the fibroblast growth factor (FGF) to its FGFR receptor (Yayon et al., 1991; Spivak et al., 1994; Venkataraman et al., 1999) and act as initial attachment receptors for bacteria (Chen,T. et al., 1995) and viruses including alphaviruses (Byrnes and Griffin, 1998), human immunodeficiency virus (HIV) type 1 (Mondor et al., 1998) and vaccinia virus (Chung et al., 1998). Heparan sulfates are components of amyloid plaques in prion diseases (Gabizon et al., 1993). We investigated the role of HSPGs as possible co-factors for 37-kDa LRP mediating PrP binding. We also constructed recombinant (rec.) SFV vectors leading to the expression of an LRP mutant termed LRPdelBD::FLAG lacking the direct-binding domain for PrP in wild-type and mutant HSPG-deficient CHO cells. We compared the PrP-binding capacity of these cells with wild-type and mutant CHO cells hyperexpressing wild-type LRP::FLAG. In light of our findings that 37-kDa LRP fails to form homodimers, and that HSPGs mediate the binding of PrP to 37-kDa LRP, the relationship between 37-kDa LRP and 67-kDa LR might be explained by the association of LRP with HSPGs as outlined in a proposed model for the LRP–PrP–HSPG complex on the cell surface.


Identification of a direct LRP interaction domain on PrP by the yeast two-hybrid system

To determine the domains of PrP interacting directly with LRP, we employed a yeast two-hybrid analysis with truncated PrP molecules in the bait position and LRP44–295 in the prey position. Only truncated PrP retaining the regions amino acids (aa) 144–179 (Figure 1A, rows 6 and 7) interacted with LRP. This region contains domains corresponding to the first α-helix (aa 144–154), the second β-strand (aa 161–164) and the first amino acid of the second α-helix (aa 179–193) of the human PrP (Zahn et al., 2000). Regions from aa 23 to 143 of the human PrP are not sufficient for binding to LRP (Figure 1A, rows 1–5). Regions from aa 180 to 230 of human PrP (row 8) are not required for the direct interaction between PrP and LRP. We termed this LRP interaction domain on PrP PrPLRPbd1.

figure cde583f1afigure cde583f1b
Fig. 1. Identification of direct PrP–LRP-interaction domains. (A) Identification of the direct PrP–LRP interaction domain on PrP. HuPrP23–93 (row 1), huPrP23–118 (row 2), huPrP23–127 (row 3), ...

Retrenchment of the direct PrP-binding domain on LRP by the yeast two-hybrid system

Recently, we mapped a direct PrP-binding domain on LRP between aa 157 and 180 (Rieger et al., 1997) employing N-terminally truncated LRP molecules. In order to re trench this binding domain precisely we co-expressed the C-terminally truncated LRP molecules LRP44–101 and LRP44–160, respectively, together with full-length PrP in the yeast two-hybrid system. Both truncations failed to interact with PrP (Figure 1B) confirming that this direct PrP-binding site coincides with the laminin-binding domain (aa 161–180). Expression of an LRP mutant lacking this direct PrP-binding domain (LRPdelBD161–180) in CHO cells (Figure 4K) showed that LRPdelBD161–180::FLAG was still able to bind to PrP, indicating the presence of a second binding site for PrP on LRP, which locates either between aa 101 and 160 or 181 and 295 of LRP.

figure cde583f4afigure cde583f4b
Fig. 4. Influence of HSPGs on the LRP–LR–PrP-binding reaction analysed by wild-type and mutant HSPG-deficient CHO cells; use of an LRP deletion mutant lacking the direct PrP-binding domain on LRP; PrP–LRP in vitro interaction studies. ...

PrP144–179 interacts directly with LRP161–179 in the yeast two-hybrid system

In order to prove a direct interaction between the PrP and the 37-kDa/67-kDa LRP/LR via PrP144–179 and LRP161– 179, we co-expressed both protein domains in bait and prey position, respectively, resulting in a strong interaction (Figure 1C, row 1). In contrast, huPrP144–179 failed to interact with LRP180–295 (row 2) or LRP44–160 (row 3).

β-galactoside gal-3 does not interact with PrP or LRP in the yeast two-hybrid system

The association of gal-3 with the LRP has been suggested (Landowski et al., 1995; Buto et al., 1998). However, gal-3 antibodies do not influence the LRP-dependent binding/internalization of PrP on the cell surface, suggesting that this molecule does not act as a co-receptor for LRP (Gauczynski et al., 2001b). For confirmation that gal-3 does not interact with PrP, we expressed gal-3 in bait and PrP in prey position of the yeast two-hybrid system resulting in no interaction between the two proteins (Figure 2A, row 2). Gal-3 also failed to interact with LRP in the yeast two-hybrid system (Figure 2A, row 3).

figure cde583f2
Fig. 2. LRP fails to interact with itself in the yeast two-hybrid system and appears monomeric by native size-exclusion chromatography. Gal-3 fails to interact with PrP and LRP. (A) huPrP23–230 and LRP fail to interact with the β-galactoside ...

37-kDa LRP fails to interact with itself in the yeast two-hybrid system and appears monomeric by size-exclusion chromatography

The polymorphism of the LR is still unclear. In order to test whether homodimerization of LRP could account for this 37-kDa/67-kDa polymorphism and to understand better the configuration of the LRP–PrP-binding complex, we cloned the cDNA encoding for LRP in the bait and prey position of the yeast two-hybrid system. LRP fails to interact with itself (Figure 2B, row 2), suggesting that LRP is unable to directly form homodimers. For confirmation we purified LRP::FLAG from SFV RNA-LRP::FLAG-transfected BHK cells by anti-FLAG antibody chromatography to homogeneity and analysed the native protein by SDS–PAGE and size-exclusion chromatography. The protein migrated as a 37-kDa protein on an SDS– polyacrylamide gel (Figure 2C, lane 1) and eluted as a 40-kDa protein from a native size-exclusion column (Figure 2D), confirming that 37-kDa LRP is monomeric under native conditions. Thus, the 67-kDa form of the LR may result from the association of the LRP with other molecules such as HSPGs.

Identification of the PrP interaction domains PrPLRPbd1 and PrPLRPbd2 by binding assays with prion peptides to NT2 and N2a cells

The yeast two-hybrid system identified the domain aa 144–179 of PrP as a direct binding site for LRP termed PrPLRPbd1. To identify other domains of PrP, which might bind indirectly to LRP, we exposed NT2 and N2a cells to glutathione S-transferase (GST)-fused PrP peptides covering the entire PrP sequence. Besides peptide 129–175 encompassing the direct binding domain PrPLRPbd1, GST::PrP53–93 bound to the cells in an LRP-dependent fashion. The binding of GST::PrP53–93 (Figure 3A) and GST::PrP129–175 (Figure 3C) is shown on NT2 cells in comparison with GST::PrP90–109 (Figure 3B). This binding can be inhibited by addition of the LRP antibody W3 (insets in Figure 3A and C). The binding properties of the whole array of peptides are also shown on N2a cells (Figure 3D–J). Only GST::PrP53–93 (Figure 3E) and GST::huPrP129–175 (Figure 3H) bound to the cells dependent on LRP–LR (LRP antibody competition is shown in the bottom insets of Figure 3). The integrin laminin receptor VLA6 does not co-localize with PrP or LRP–LR on the surface of neuroblastoma cells (Gauczynski et al., 2001b). The addition of an anti-VLA6 antibody, failed also to compete for the binding of GST::PrP53–93 or GST::PrP129–175 to N2a cells (Figure 3E and H, top insets), confirming that VLA6 does not act as a receptor for PrP. We termed the indirect binding domain PrPLRPbd2. As the two binding domains are located N- and C-terminally of the proteinase K cleavage site of PrPres, we tested longer peptides in our binding assay corresponding to the two fragments that result from proteolytic cleavage of PrP, i.e. PrP23–89 and PrP90–230. Both peptides bound to both cell types in an LRP-dependent manner (Table (TableI).I). Combining these data with the results from the yeast two-hybrid system (Figure 1) we conclude that two binding sites on PrP for LRP termed PrPLRPbd1 (aa 144–179) and PrPLRPbd2 (aa 53–93) do exist. Results of the PrP peptide binding studies to N2a and NT2 cells including antibody competitions are summarized in Table TableII.

figure cde583f3
Fig. 3. Identification of PrP-interaction domains for LRP/LR by binding assays with rec. prion peptides on NT2 and N2a cells. (A–C) Binding to NT2 cells. NT2 cells were incubated with PrP peptides fused to GST in the absence (A–C) ...
Table I.
Summary of the binding behaviour of individual GST-fused PrP peptides to NT2 and N2a cells including LRP–LR and VLA6 antibody competition

Binding of PrP to LRP via PrPLRPbd2 is dependent on HSPGs

The cell-binding assay led to the identification of an additional binding domain for LRP on PrP which was not identified in the yeast two-hybrid system, indicating that a third molecule is necessary to mediate the binding of LRP to PrPLRPbd2. It has been reported that 60% of the binding of rec. chicken PrP to CHO cells depends on the presence of endogenous heparan sulfates (Shyng et al., 1994). Mutant CHO cells (S745) are severely deficient in HSPGs because of an altered xylose transferase activity, the first enzyme required for glycosylaminoglycan (GAG) synthesis (Esko et al., 1985) and, therefore, represent an appropriate model system to investigate whether HSPGs might represent the third interactor in the binding of PrPLRPbd2 to LRP. First, we proved that the binding of rec. GST::huPrP23–230 to wild-type CHO cells (Figure 4A) and to the mutant CHO cells (Figure 4B) was LRP–LR-dependent (Figure 4C and D). We then saturated selectively PrPLRPbd1 by incubating GST::huPrP23–230 with a monoclonal PrP antibody directed against the domain 140–180 of PrP. Obstructing PrPLRPbd1 inhibited the binding of the rec. PrP to HSPG-deficient (Figure 4F) but not to wild-type CHO cells (Figure 4E), demonstrating that binding of PrP to LRP via PrPLRPbd2 needs the presence of HSPGs. The peptide GST::PrP53–93 corresponding to PrPLRPbd2, which bound to normal cells (Figure 4G), did not bind to HSPG-deficient cells (Figure 4H). However, the binding was restored in a dose-dependent manner after addition of soluble HSPGs (Figure 4I and J), confirming that the interaction of PrP to LRP via PrPLRPbd2 is HSPG dependent.

Identification of an HSPG-dependent second binding site for PrP on LRP

The two binding sites on PrP, one of which is direct and the other indirect, suggested that two ‘acceptor’ sites might also exist on LRP. To test this hypothesis we adapted our SFV expression system to CHO cells. We then expressed LRP::FLAG and a mutant LRP lacking the direct PrP-binding domain (aa 161–180), termed LRPdelBD::FLAG, in wild-type CHO cells and the mutant CHO-S745 cell line lacking HSPGs (Figure 4K, lower panels). In wild-type CHO cells (Figure 4K, lanes 1–6) the binding of GST::huPrP was enhanced, when LRP::FLAG was hyperexpressed (Figure 4K, lanes 2 versus 6). Hyper expression of LRPdelBD::FLAG did not reduce the amount of the bound GST::huPrP (Figure 4K, lane 4), indicating that the binding was HSPG mediated. In mutant CHO-S745 cells lacking HSPGs (Figure 4K, lanes 7–13) the amount of bound GST::huPrP was also enhanced in LRP::FLAG hyperexpressing cells compared with non-transfected cells (Figure 4K, lanes 8 and 13). CHO-S745 cells expressing LRPdelBD::FLAG, however, showed a reduced binding of GST::huPrP (Figure 4K, lane 10), similar to non-transfected cells (Figure 4K, lane 13) suggesting that the second indirect PrP-binding site was not functioning in the absence of HSPGs. In order to confirm that HSPGs are responsible for the binding of PrP to the second binding domain on LRP, we added HSPGs to cells overexpressing LRPdelBD::FLAG resulting in a total restoration of the PrP binding (Figure 4K, lane 11). HSPGs failed to increase the binding of GST::huPrP to CHO wild-type (Figure 4L, lanes 2 and 3) and CHO-S745 cells (Figure 4L, lanes 5 and 6) due to the presence of the direct binding domains on PrP and LRP. We conclude from these data that two binding sites for PrP on LRP exist: a direct one, which is located from aa 161–179, and a second indirect one, which resides either between aa 101 and 160 or between aa 180 and 295 of LRP.

Interaction of PrP and LRP in vitro

The direct binding domains on LRP (aa 161–179) and PrP (aa 144–179) should allow the two proteins to interact with each other in vitro. GST-fused LRP (Rieger et al., 1997) and immobilized FLAG::huPrP (Figure 4M, lanes 1 and 2) were able to interact with each other in vitro as shown in the pull down assay depicted in Figure 4M. As already observed on wild-type CHO and CHO-S745 cells, HSPGs did not influence the interaction due to the presence of the direct interaction domains (Figure 4M, lanes 2 and 3). However, HSPGs did affect the LRP–PrP53–93 interaction (HSPG-dependent binding domain on PrP) and the LRPdelBD–PrP interaction (lacking the direct binding domain on LRP) in CHO-S745 cells (Figure 4G–J and K, respectively).


Cell-binding and internalization studies proved that the 37-kDa LRP/67-kDa LR acts as the receptor for the cellular PrP, PrPc, on the cell surface (Gauczynski et al., 2001b). In order to investigate the interaction domains on PrP and LRP–LR mediating the binding of these two proteins a series of interaction studies employing the yeast two-hybrid system as well as PrP-binding assays with neuronal and non-neuronal cells including the SFV system have been performed.

Mapping of LRP interaction sites on PrP

First, we aimed to determine which part of the PrP interacts with LRP. We performed a yeast two-hybrid analysis with a series of PrP deletion variants and employed a series of PrP peptides covering the entire PrP in various cell-binding assays.

In the yeast two-hybrid system, we identified the PrP domain aa 144–179 as a direct LRP-binding domain, termed PrPLRPbd1. This binding domain was confirmed in cell-binding assays in N2a and NT2 cells with PrP peptides encompassing the entire PrP sequence. As already observed with full-length PrP (Gauczynski et al., 2001b), the staining pattern with the PrP peptides to N2a and NT2 cells is also punctuate due to either receptor clustering or PrP–PrP peptide aggregation, or both. Receptor clustering was observed with a variety of other cell-surface receptors such as the FGF receptor (Utton et al., 2001), the muscle nicotinic acetylcholine receptor (AChR) (Hoch et al., 2001) or the tumour necrosis factor receptor (TNFR55) (De Wilde et al., 2001). In addition to the yeast two-hybrid assay, which identified the direct PrP–LRP–LR interaction domain PrPLRPbd1, the cell-binding assay identified a second binding domain between aa 53 and 93 of PrP, termed PrPLRPbd2. This domain was not functional in the yeast two-hybrid system, indicating that an additional factor lacking in the yeast cell nucleus is required for mediating the interaction between LRP and PrP through PrPLRPbd2. Employing a mutant HSPG-deficient CHO cell line, the binding assay revealed that PrPLRPbd2 binds to LRP via HSPGs. This finding was confirmed by using the PrP peptide 53–93 corresponding to PrPLRPbd2. PrP53–93 failed to bind to the mutant CHO cells but the binding was restored in a dose-dependent manner by the addition of soluble HSPGs.

Our mapping data, which resulted in the identification of two LRP-binding sites on PrP, are consistent with the results obtained with PrP knock-out mice expressing PrPs with N-proximal deletions, suggesting that PrP could bind to its natural ligand, termed Lprp, which could represent a PrP receptor, in a region C-terminal to aa 134 of the PrP (Shmerling et al., 1998). In line with this study, PrPLRPbd1 would correspond to the Lprp-binding domain. The same authors hypothesized the existence of a second domain located more N-terminally initiating a signal transduction necessary to maintain normal cellular functions. Recently, a signal transduction activity of the PrP by activation of the tyrosine kinase Fyn was described (Mouillet-Richard, 2000). As PrP resides as a GPI-anchored protein outside the cell, whereas the tyrosine kinase Fyn locates to the inner plasma membrane inside the cell, transmembrane orientated LRP–LR might be a reasonable candidate mediating the intracellular signal transduction between PrPc and Fyn. The cytosolic domain of N-syndecans (syndecan-3), one of a family of four transmembrane cell surface HSPGs, has recently been shown to bind to complexes containing c-Src and Fyn kinases (Kinnunen et al., 1998). This activity is related to neurite outgrowth (Kinnunen et al., 1998). Hence, it is also conceivable that HSPGs might be necessary for the so far hypothetical LRP–LR-mediated signal transduction between PrPc and Fyn. A possible intercellular role of the PrP–LRP–LR interaction involving HSPGs which might result in signalling or cell attachment (for review see Gauczynski et al., 2001a) will be further investigated in detailed cell–cell interaction assays.

Recently, studies on the transmission of human prions to transgenic mice suggested that a so far unidentified protein X may participate in the formation of PrPres. Substitution of residues 167, 171 and 218 prevented PrPres formation suggesting that protein X may interact with those residues of the PrP (Kaneko et al., 1997). As aa 167 and 171 reside within the direct PrP–LRP interaction domain termed PrPLRPbd1 it cannot be excluded that LRP–LR might function as protein X. Application of LRP–LR in PrP oligo/multimerization processes may enlighten a possible role of the 37-kDa/67-kDa LR in PrPres formation.

Mapping of the PrP interaction sites on LRP

A yeast two-hybrid analysis with C-terminal LRP truncations extended a previous analysis (Rieger et al., 1997) and retrenched the direct PrP-binding domain to aa 161–179 of LRP. As the binding domain PrPLRPbd2 on PrP is HSPG dependent, we hypothesized that a second indirect binding domain for PrP may also exist on LRP. To test this hypothesis, we analysed PrP-binding to wild-type CHO cells, to mutant HSPG-deficient CHO cells and to both cell types hyperexpressing either the full-length LRP::FLAG or an LRP mutant lacking the direct PrP-binding domain (aa 161–180), termed LRPdelBD::FLAG. HSPGs failed to increase the binding of GST::huPrP to CHO wild-type and CHO-S745 cells due to the presence of the direct binding domains on PrP and LRP. CHO and CHO-S745 cells transfected with recombinant SFV RNAs further revealed the role of HSPGs in the binding of PrP to LRP. Hyperexpression of LRP::FLAG or LRPdelBD::FLAG in wild-type CHO cells resulted in approximately the same increase of GST::huPrP-binding when compared with non-transfected cells. This indicates that binding of PrP to LRP::FLAG can occur via a second PrP-interaction domain present in LRPdelBD::FLAG. In HSPG-deficient CHO cells, however, PrP-binding to cells hyperexpressing LRPdelBD::FLAG was similar to non-transfected cells, indicating that the second interaction domain on LRP was not functioning. In contrast, LRP::FLAG hyperexpressing HSPG-deficient CHO cells showed an increased binding of GST::huPrP due to the presence of the direct HSPG-independent PrP-binding domain (aa 161–179). The addition of HSPGs to LRPdelBD::FLAG hyperexpressing HSPG-deficient CHO cells restored GST::huPrP binding to levels achieved with LRP::FLAG hyperexpressing cells. The data demonstrate the existence of a second HSPG-dependent PrP-binding domain on LRP, residing either between aa 101 and 160 or aa 181 and 285 of LRP. The identification of a HSPG-binding domain on LRP–LR which resides between aa 205 and 229 of LRP–LR (Kazmin et al., 2000) suggests that this domain may represent the HSPG-dependent-binding site for PrP. A monoclonal antibody directed against aa 167–243 of LRP reduces PrP-binding to neuronal cells (Gauczynski et al., 2001b) suggesting that the indirect binding domain may reside between aa 180 and 285 rather than aa 101–160 of LRP.

Interaction of PrP and LRP in vitro

Rec. purified PrP and LRP interact with each other in vitro due to the presence of the direct PrP–LRP interaction domains. The influence of HSPGs on the indirect PrP–LRP interaction domains, however, was only detectable on the HSPG-deficient CHO-S745 cells employing PrP53–93 (representing the indirect binding domain on PrP) and LRPdelBD (lacking the direct binding domain on LRP).

The relationship between 37-kDa LRP and 67-kDa LR

Attempts to isolate the gene for the 67-kDa LR revealed a cDNA fragment encoding the 37-kDa LRP (Rao et al., 1983; Yow et al., 1988; Grosso et al., 1991). Pulse–chase experiments with 37-kDa LRP specific antibodies demonstrated that 37-kDa LRP is the precursor of 67-kDa LR (Rao et al., 1989; Castronovo et al., 1991). The 37-kDa molecule encoded by the full-lengh gene identified for the LR, is virtually identical to the ribosomal protein p40. The 37LRP/p40 evolved from a ribosomal protein essential for protein synthesis lacking any laminin-binding abilities, to a laminin-binding cell surface receptor (for review see Ardini et al., 1998). The molecular structure of the 37-kDa LRP/67-kDa LR and the mechanism by which the 37-kDa LRP forms the mature 67-kDa LR remains unclear. Our data from a yeast two-hybrid analysis show that LRP fails to interact with itself, an argument against the hypothesis of a direct homodimerization. In addition, homogeneous rec. LRP::FLAG appears to be monomeric as analysed by size-exclusion chromatography. Although we only used 0.1% Triton X-100 in the purification procedure, we cannot exclude that this small amount of non-ionic detergent may disrupt a native dimeric state of the receptor. A recent study suggested that acylation of LRP would allow it to associate with a heterologous molecule (Buto et al., 1998). In the light of our results, the relationship between the 37-kDa LRP and 67-kDa LR might be explained by the association of an LRP molecule with heparan sulfates.

Role of heparan sulfates in the PrP–LRP interaction process

The binding of chicken PrP to the surface of mammalian cells has been shown to depend partly on the availability of heparan sulfates expressed by these cells (Shyng et al., 1995). PrP interacts with heparan sulfates (Gabizon et al., 1993; Caughey et al., 1994; Chen,S.G. et al., 1995; Brimacombe et al., 1999). Recent studies demonstrated that the binding of copper to PrP which occurs within the octarepeat region (Brown et al., 1997) (corresponding to PrPLRPbd2) can be competed by the addition of HSPGs (Brimacombe et al., 1999), confirming that this region of PrP binds to HSPGs. Finally, LRP–LR has also been shown to be a heparin/heparan sulfate-binding molecule (Guo et al., 1992; Kazmin et al., 2000). The predicted α-helical structure (aa 205–229) of LRP–LR is proposed to have heparin-binding characteristics (Kazmin et al., 2000). The requirement of heparin-like molecules for the formation of a ligand–receptor complex is not unprecedented and is well illustrated by the example of the binding of the FGF to its receptor, FGFR (Yayon et al., 1991; Spivak et al., 1994; Venkataraman et al., 1999). All these data match very well with our findings, providing a comprehensive model of the PrP–LRP–LR interaction (Figure 5). A heparan sulfate arm of a cell-surface HSPG molecule might be located between PrPLRPbd2 and the indirect binding domain of LRP to create a sandwich interaction site, whereas PrP would inter act with LRP–LR (aa 161–179) directly via PrPLRPbd1. We cannot exclude that HSPGs intervene in a more indirect manner, by changing the conformation of LRP–LR as to render it amenable to its interaction with the PrPLRPbd2 region of PrP.

figure cde583f5
Fig. 5. Model for the function of LRP–LR as the receptor for PrP. The PrP molecule binds to LRP–LR via PrPLRPbd1 and PrPLRPbd2. PrPLRPbd2 (aa 53–93) is dependent on the presence of a heparan sulfate arm of a HSPG molecule whereas ...

Components blocking the direct and indirect PrP–LRP interaction domains on PrP and LRP may represent a novel class of molecules suitable for a therapeutic intervention in prion diseases.

Materials and methods

Construction of pSFV1-LRPdelBD::FLAG-preparation of SFV mRNAs in vitro

The LRP mutant pSFV1-LRPdelBD::FLAG (deletion of the direct PrP-binding domain located between aa 161 and 180) was generated by the QuikChange™ site-directed mutagenesis method (Stratagene) using the pSFV1-LRP::FLAG plasmid DNA (Gauczynski et al., 2001b) as template for PCR and confirmed by dideoxy sequencing. The rec. plasmid DNAs pSFV3-lacZ (Life Technologies), pSFV1-LRP::FLAG and pSFV1-LRPdelBD::FLAG were linearized, purified and transcribed as described (Gauczynski et al., 2001b). The correct length of the transcripts was verified by agarose gel electrophoresis. RNA was stored at –20°C.

Mammalian cell culture, transfection and co-transfection studies with the SFV system

Mutant CHO cells (S745) deficient in xylose transferase (Esko et al., 1985) as well as wild-type CHO (K1) were cultivated in NUT.MIX.F-12(HAM) supplemented with GLUTAMAX-I (Gibco-BRL), 10% fetal calf serum (FCS), 100 µg/ml penicillin and 100 µg/ml streptomycin at 37°C with 5% CO2. Transfections and co-transfections were carried out as described (Gauczynski et al., 2001b). Transfection efficiencies as determined by transfecting SFV3-lacZ control RNA followed by X-gal staining were ~80% for CHO-K1 or CHO-S745 cells.

Purification of LRP::FLAG from the SFV system

Transfection of BHK cells with the rec. SFV LRP::FLAG RNA was performed as described (Gauczynski et al., 2001b). The total volume of the electroporated cells was plated on 10-cm dishes containing 15 ml of complete growth medium followed by incubation for at least 48 h at 37°C. Forty-eight hours post-transfection, cells were harvested, washed once with phosphate-buffered saline (PBS) and then lysed in PBS supplemented with 0.1% Triton X-100 by repeated freezing and thawing. The crude lysate was obtained by centrifugation at 14 000 r.p.m., 4°C for 15 min and purified by the batch method using an anti-FLAG M2 affinity gel (Sigma). The FLAG-tagged protein was bound over night by rotating at 4°C, washed four times with Tris-buffered saline (TBS), eluted over night by competition with 1 ml TBS containing 100 µg/ml FLAG peptides and dialysed against 20 mM HEPES pH 7.4. The purity and the concentration of the protein was checked by SDS–PAGE followed by silver staining of the gel.

Recombinant proteins generated in the Baculovirus and Escherichia coli system

Rec. GST, GST::huPrP23–230, GST::haPrP23–89 and GST::haPrP90– 231 were expressed in Baculovirus infected Sf9 cells and purified to homogeneity as described for hamster GST::PrP fusions previously (Weiss et al., 1995, 1996). GST-fused PrP-peptides (haPrP23–52, haPrP53–93, haPrP90–109, haPrP129–175, haPrP180–210, haPrP218– 231) were expressed in Escherichia coli and purified to homogeneity as described for GST fusions (Weiss et al., 1995). cDNA encoding for huPrP110–128 was cloned via EcoRI (5′) and BamHI (3′) into pGEX-2T, GST::huPrP110–128 was expressed in E.coli and purified to homogeneity as described (Weiss et al., 1995). All rec. proteins were dialysed against 20 mM HEPES, pH 7.4.

PrP-binding assays followed by immunofluorescence analysis

N2a, and human NT2 cells were maintained in DMEM medium con taining 10% FCS, 1% glutamine, 100 µg/ml penicillin and 100 µg/ml streptomycin. Mutant CHO cells (S745) and wild-type CHO-K1 were cultivated as described above. For competition studies, the cells were either pre-incubated for 2 h with the individual antibody diluted in culture medium or co-incubated with rec. protein and antibody (inoculum saturation). In case of pre-incubation, medium was replaced and cells were incubated overnight with 4 µg/ml of rec. GST-fusion proteins per ml of culture medium. Cells were then washed three times with PBS and prepared for immunofluorescence microscopy, which was performed as described (Gauczynski et al., 2001b).

PrP-binding assay in cell culture followed by western blotting

CHO/CHO-S745 cells (8 × 105) (either non-transfected or transfected with rec. SFV RNAs) were seeded on 6-well plates and incubated at 37°C. Twenty-four hours post-transfection, cells were incubated in medium containing 5 µg/ml of rec. GST–huPrP23–230 for 18 h at 37°C. Together with the rec. protein, cells were co-incubated with 40 µg/ml of HSPGs (when indicated). Cells were then washed several times with PBS and scraped off in PBS. After centrifugation the pellets were resuspended in lysis buffer (25 mM Tris–HCl pH 7.4, 150 mM NaCl, 1 mM CaCl2, 3 mM MgCl2, 1% NP-40). After addition of Laemmli buffer, samples were separated by SDS–PAGE and blotted on PVDF membrane. Western blotting was performed with the monoclonal anti-PrP antibody 3B5 or the pAb LRP W3 and peroxidase-coupled secondary antibodies.

Mapping of LRP and PrP-binding sites in the yeast two-hybrid system

Constructions of plasmids pSH2-1 and pJG4-5 were described previously (Rieger et al., 1997). For mapping the LRP–PrP interaction site on PrP, the following C-terminal truncated constructs of PrP were generated: pSH2-1-GST::huPrP23–93, pSH2-1-GST::huPrP23–118, pSH2-1-GST:: huPrP23–127, pSH2-1-GST::huPrP23–131, pSH2-1-GST::huPrP23–143, pSH2-1-GST::huPrP23–154, pSH2-1-GST::huPrP23–181 and pSH2-1-GST::huPrP180–230. The PrP fragments were amplified by PCR using oligodesoxyribonucleotides coding for different PrP sequences flanked by a BamHI (5′) and a SalI (3′) restriction site. The fragments were cloned via BamHI and SalI into the vector pSH2-1-GST. All PrP constructs were confirmed by sequencing. The different bait plasmids, the prey plasmid pJG4-5-LRP44–295 and the reporter plasmid pSH18-34 (lacZ) were co-transformed into EGY48 cells and transformants were tested in β-galactosidase assays. Construction of the plasmid pSH2-1-GST::huPrP23–230 was described previously (Rieger et al., 1997). For mapping the PrP interaction site on LRP, the C-terminal truncated constructs of LRP pJG4-5-LRP44–101 and pJG4-5-LRP44–160 were designed. The LRP fragments were amplified by PCR using oligodesoxyribonucleotides coding for different LRP sequences flanked by EcoRI (5′) and SalI (3′). The fragments were cloned via EcoRI and XhoI restriction sites into the vector pJG4-5. The resulting constructs were confirmed by didesoxysequencing. The different bait plasmids, the prey plasmids and the reporter plasmid pSH18-34 (lacZ) were co-transformed into EGY48 cells and transformants were tested in a β-galactosidase assays.

LRP–LRP and gal-3–PrP interaction studies in the yeast two-hybrid system

The constructs pSH2-1-GST, pJG4-5-GST, pSH2-1-GST::huPrP and pJG4-5-LRP44–295 were described previously (Rieger et al., 1997). The LRP44–295 encoding cDNA was PCR amplified from pJG4-5-LRP44–295. The PCR product flanked by EcoRI (5′) and SalI (3′) restriction sites, respectively, was cloned into the vector pSH2-1 via both restriction sites resulting in pSH2-1-LRP44–295, which was confirmed by dideoxy sequencing. Yeast transformations and dotting were carried out as described above. Total RNA was isolated from 293 cells by RNeasy kit (Qiagen). Gal-3 cDNA was amplified by RT–PCR and subcloned into pSH2-1 via EcoRI (5′) and SalI (3′) restriction sites resulting in pSH2-1-Gal-3, which was confirmed by dideoxy sequencing. The huPrP23–230 encoding cDNA was excised from pSH2-1-GST::huPrP23–230 and subcloned into pJG4-5 via EcoRI and SalI. The resulting plasmid pJG4-5-GST::huPrP23–230 was confirmed by didesoxy sequencing. Yeast transformations and dotting were carried out as described above.

Analysis of native LRP::FLAG by size-exclusion chromatography

The Superose 12 PC 3.2/30 column (Amersham Pharmacia) was calibrated with the LMW calibration kit in 20 mM HEPES pH 7.4. Purified LRP-FLAG (2.5 µg) expressed in the SFV system was loaded in a total volume of 25 µl. Chromatography was performed at a flow rate of 30 µl/min. The eluted LRP was detected with a UV-M II monitor at 280 nm.

In vitro interaction of PrP and LRP

Rec. FLAG::huPrP23–230 was expressed in the Baculovirus system (C.Hundt et al., manuscript in preparation) according to rec. FLAG::haPrP23–231 (Rieger et al., 1997) and immobilized on anti-FLAG M1 beads. Twenty microlitres of beads (1:1 slurry; 500 ng of FLAG::huPrP), were incubated in TBS supplemented with 2 mM MgCl2 with 500 ng of rec. GST and 1 µg of GST::LRP (Rieger et al., 1997), respectively (molar ratio: 1:1) in the absence and presence of 1.5 µg HSPGs (Sigma). Homogeneity of GST::LRP, GST and FLAG:: huPrP23–231 was proven on silver-stained SDS–PA gels. After 1 h at room temperature, the supernatant was removed, beads washed four times with TBS, boiled in SDS-sample buffer and analysed by western blotting developed with mAb GST.


pAb LRP W3 (Rieger et al., 1997) was purified by protein A–Sepharose chromatography. mAb GST (Santa Cruz Biotechnology), mAb VLA6 (Immunotech.), mAb 3B5 (G.Hunsmann), mAb SAF70 (aa 140–180 of PrP) and pAb JB007 (CEA, France), secondary fluorescein isothiocyanate (FITC), Cy3 (indocarbocyanine) and Texas Red-conjugated antibodies (used at 1:100 dilutions; Jackson Laboratories/Southern Biotechnology) were used.


We thank D.Barritault (Université Paris XII, Créteil, France) for delivering CHO-S745 cells, J.Grassi (CEA-Saclay, France) for the SAF70 antibody, G.Hunsmann for the 3B5 antibody and J.Turnbull (MRC, Birmingham) for helpful discussions and critical reading of the manuscript. We thank S.Janetzky, S.Hengge, A.Pahlich and K.Krüger (Munich) for excellent technical assistance. This work was supported by grant from DRET (Direction des Recherches et Techniques, Paris, France), INSERM (Paris, France) and from the European Union (BIOMED PL-97-6054, QLRT-2000-02085 and FAIR-CT-98-7020). S.W. acknowledges the support of grants KI-01-9760 and KO-01-1541-01 (BMBF), LMU 3 and 4 (Bavarian Prion Research Foundation) and thanks R.Grosschedl and E.-L.Winnacker for valuable advice and continuous support.


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