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Originally published In Press as doi:10.1074/jbc.M400952200 on March 18, 2004

J. Biol. Chem., Vol. 279, Issue 24, 25058-25065, June 11, 2004
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Characterization of Recombinant, Membrane-attached Full-length Prion Protein*

Heike Eberl{ddagger}, Peter Tittmann§, and Rudi Glockshuber{ddagger}

From the {ddagger}Institut für Molekularbiologie und Biophysik, Eidgenössische Technische Hochschule Hönggerberg and §Elektronenmikroskopie-Zentrum der ETH Zürich, incare Institut für Angewandte Physik, Eidgenössische Technische Hochschule Hönggerberg, CH-8093 Zürich, Switzerland

Received for publication, January 28, 2004 , and in revised form, March 16, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
An abnormal isoform, PrPSc, of the normal cellular prion protein (PrPC) is the major component of the causative agent of prion diseases. Both isoforms were found to possess the same covalent structures, including a C-terminal glycosylphosphatidylinositol anchor, but different secondary and tertiary structures. In this study, a variant of full-length PrP with an unpaired cysteine at the C terminus was recombinantly produced in Escherichia coli, covalently coupled to a thiol-reactive phospholipid, and incorporated into liposomes to serve as a model for studying possible changes in structure and stability of recombinant PrP upon membrane attachment. Covalent coupling of PrP to liposomes did not result in significant structural changes observable by far-UV circular dichroism. Moreover, limited proteolysis experiments failed to detect changes in the stability of liposome-bound PrP relative to soluble PrP. These data suggest that the requirement of raft localization for the PrPC to PrPSc conversion, observed previously in cell culture models, is not because of a direct influence of raft lipids on the structure and stability of membranebound PrPC but caused by other factors, e.g. increased local PrP concentrations or high effective concentrations of membrane-associated conversion factors. The availability of recombinant PrP covalently attached to liposomes provides the basis for systematic in vitro conversion assays with recombinant PrP on the surface of membranes. In addition, our results indicate that the three-dimensional structure of mammalian PrPC in membranes is identical to that of recombinant PrP in solution.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Transmissible spongiform encephalopathies (TSEs)1 are fatal neurodegenerative disorders that affect humans and other mammalian species. The "protein-only" hypothesis states that the causative agent of all TSEs, the prion, is devoid of informational nucleic acids and consists mainly, if not entirely, of PrPSc, an abnormal, oligomeric, and proteinase-resistant isoform of the monomeric, host-encoded, cellular prion protein PrPC (reviewed in Refs. 1–4). Although there are significant differences in the biophysical properties of PrPC and PrPSc (5–11), no differences in the covalent structures of PrPC and PrPSc could be detected so far (12–14). Both isoforms harbor a single disulfide bond (15), consist of mixtures of non-, mono-, or diglycosylated PrP forms, and possess a C-terminal glycosylphosphatidylinositol (GPI) anchor (16).

Several lines of evidence imply that the C-terminal GPI anchor might play a role in the conversion of PrPC to PrPSc. PrPC was found to be susceptible to GPI anchor cleavage by phospholipase C, whereas PrPSc under the same conditions was not susceptible (17, 18). Furthermore, the GPI anchor directs the prion protein to certain microdomains in the cell membrane, so-called rafts that are rich in cholesterol and lipids with saturated acyl chains, mainly phosphatidylcholine lipids and sphingomyelin (19–22). This raft localization was found to be essential for the formation of PrPSc in vivo. Experiments with prion proteins in which the GPI anchor was replaced by different C-terminal transmembrane helices, ensuring PrP localization on the cell surface but preventing the sorting of PRP to rafts, showed that proteinase K resistance, a hallmark of PrPSc, could not be propagated from these PrP variants (23, 24). Moreover, depletion of cholesterol from cellular membranes, known to lead to a disruption of raft microdomains (25), also prevented PrPSc formation (24), whereas sphingolipid depletion was found to increase PrPSc formation in prion-infected cell cultures (26). Further in vitro conversion experiments with raft preparations from prion-infected and non-infected cells, as well as raft-like liposomes, demonstrated that PrPC and PrPSc must either be co-localized in the same membrane or that PrPC must be released by GPI anchor cleavage for conversion to occur (27, 28). Although these results strongly suggested a direct influence of the covalent membrane attachment of the prion protein on the conversion process, previous studies (29–31) on the influence of a membranous environment on the structure and stability of PrPC were performed with soluble, recombinant prion proteins lacking a membrane anchor.

In the present study, we constructed a variant of the recombinant murine prion protein that can be attached covalently to membranes. The variant, termed PrP-SH, consists of the sequence of the full-length prion protein, mPrP-(23–231), with a six amino acid extension (-Gly-Gly-Gly-Gly-Gly-Cys-CO2–) at the C terminus. The C-terminal cysteine residue of the extension contains the only thiol group in PrP-SH and can be covalently coupled to a thiol-reactive lipid (PDP-DHPE) incorporated in preformed liposomes (Fig. 1). Lipid-coupled PrP-SH was termed PrP-SS-PDHPE and allowed, for the first time, the comparison of soluble, recombinant PrP with recombinant PrP covalently incorporated into liposomes. Our data demonstrate that covalent coupling of PrP to cellular membranes does not significantly influence the overall structure of recombinant PrP, indicating that the NMR structures solved for soluble, recombinant PrPs from mouse, hamster, man, and cattle (32–39) in the absence of membranes most likely represent the non-liganded conformation adopted by PrPC on the cell surface. Moreover, synthetic liposomes bearing PrP-SS-PDHPE provide the basis for future in vitro conversion experiments with membrane-bound, recombinant PrP.



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  		FIG. 1.
Schematic representation of PrP-SS-PDHPE, synthesized from PrP-SH by covalent coupling to PDP-DHPE; the unstructured N-terminal segment 23–120 of PrP-(23–231) is indicated by a dotted line; the tertiary structure of the C-terminal 121–231 domain is shown as ribbon representation, and the C-terminal linker sequence of PrP-SH is given in the three-letter code.

 

   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Expression Plasmid Construction—First, a C-terminal six-amino acid linker consisting of five glycine residues and one cysteine residue was fused to the genetic sequence of the murine full-length prion protein. The product of a PCR with the plasmid pPrP-(23–231) (40) as template and the primers N1, 5'-GAC TGA TGT CCA TAT GTC TAA AAA GCG TCC AAA GCC TGG AGG GTG GAA CAC CG-3', and C1, 5'-TTC GGA TCC ATT ACT AAC ACC CCC CCC CCC CCC CGC TAG AAC GAC GCC CAT CGA ATA-3', was ligated into the pRBI-PDI-T7t-NdeI plasmid (41) via the NdeI and BamHI restriction sites, resulting in the plasmid pPrP-SH. Next, pPrP-SH was used as template for a PCR with the following primers, N2 5'-CGG GTG GAT CCA AAA AGC GGC CAA AGC CTG GAG GG-3' and C2 5'-AGC TTC GAA TTC TTA CTA ACA CCC CCC CCC CCC CCC-3', and cloned into pRSETA (Invitrogen) (42) via the BamHI and EcoRI restriction sites, which added a thrombin-cleavable His6 tag to the N terminus of the PrP-SH sequence. The new plasmid was termed pRSETA(PrP-SH).

Expression and Purification of PrP-SH and PrP-(23–231)—Production of PrP-SH was essentially based on the procedure described by Zahn et al. (62). Cells of Escherichia coli BL21 (DE3) harboring pRSETA(PrP-SH) for cytoplasmic expression of N-terminally His6-tagged PrP-SH were grown at 37 °C in 10 liters of LB medium containing 100 µg/ml ampicillin. When an optical density at 600 nm (OD600) of 0.6 was reached, protein expression was induced by addition of isopropyl-1-thio-{beta}-D-galactopyranoside to a final concentration of 1 mM. Cells were grown further for another 8 h, harvested by centrifugation, suspended in 200 ml of 50 mM Tris/HCl, pH 8.0, 1 mM MgCl2, containing 5 mg/liter LB DNase, 5 mg/liter LB RNase, 15 mg/liter LB lysozyme, and 1 tablet of protease inhibitor mixture (Roche Applied Science), and disrupted at 4 °C by sonication. The lysate was centrifuged, and sedimented His6-PrP-SH inclusion bodies were washed three times with washing buffer (20 mM Tris/HCl, pH 8.0, 23% (w/v) sucrose, 0.5% (v/v) Triton X-100, 1 mM benzamidine) until the supernatant stayed clear. Inclusion bodies were then dissolved in 6 M GdmCl, 10 mM Tris/HCl, pH 8.0, 100 mM sodium phosphate, 10 mM reduced glutathione. After another centrifugation step, the supernatant was applied to a Ni-NTA column equilibrated with the same buffer. The column was washed with 4 volumes of equilibration buffer before a linear gradient to 10 mM Tris/HCl, pH 8.0, 100 mM sodium phosphate was applied to allow oxidative on-column refolding of His6-PrP-SH. Protein impurities were removed by washing with 2 column volumes of 50 mM imidazole/HCl, 10 mM Tris/HCl, pH 8.0, 100 mM sodium phosphate. His6-PrP-SH was then eluted with 10 mM Tris/HCl, pH 5.8, 100 mM sodium phosphate, 500 mM imidazole/HCl. The eluate was extensively dialyzed against water. The His6 tag was then removed by thrombin cleavage for 1 h at room temperature in 5 mM Tris/HCl, pH 8.6, using 0.1 unit thrombin/ml His6-PrP-SH solution (~30 µM). In order to remove thrombin and artificial disulfide-linked dimers of PrP-SH, the protein solution was dialyzed against 10 mM NaH2PO4/NaOH, pH 6.5, 10 mM dithiothreitol, i.e. conditions under which the structural disulfide bond of the prion protein is resistant against reduction, but disulfide-linked dimers are reduced and applied to a CM52 cation exchange column. Elution of PrP-SH was achieved by a linear NaCl gradient from 0 to 500 mM NaCl. Fractions containing pure PrP-SH were combined, dialyzed against water, and stored at –20 °C until further use. The correct mass of PrP-SH was confirmed by MALDI-TOF mass spectrometry (calculated mass, 23,551 Da; measured mass, 23,556 Da). Complete cleavage of the His6 tag was further shown by N-terminal Edman sequencing, and the presence of one accessible thiol per PrP-SH was verified by Ellman's assay (42). The final yield of purified PrP-SH was 5 mg/liter bacterial culture. Purification of PrP-(23–231) was performed as described previously (43).

Protein Concentrations—The protein concentrations were calculated according to Gill and von Hippel (63) using the specific absorbances of the proteins. Values (A280nm, 1 mg/ml, 1 cm) of 2.70 for mPrP-(23–231) (40) and 2.66 for PrP-SH.

Preparation of Liposomes—All lipids for the preparation of liposomes were purchased from Sigma. Two kinds of liposomes differing in lipid composition were prepared. The first species, termed PC/Chol liposomes, were composed of phosphatidylcholine and cholesterol in a molar ratio of 1:1. The second species, termed raft liposomes, consisted of phosphatidylcholine, phosphatidylethanolamine, sphingomyelin, cerebrosides, and cholesterol in a molar ratio of 1:1:1:1:2 (20). This lipid composition mimicked the composition of detergent-insoluble microdomains in the cell membrane (rafts) (44, 45). Both liposome species also contained 1 mol % of a thiol-reactive lipid, PDP-DHPE (N-((2-pyridyldithio)propionyl)-1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine, triethylammonium salt) (Molecular Probes), for covalent coupling of PrP-SH to the liposomes via thiol/disulfide chemistry. Solutions of the respective lipid compositions in chloroform were dried in a rotary evaporator until the lipids formed a thin layer inside the flasks. The lipid films were suspended in water at a total lipid concentration of 10 mg/ml. The resulting milky suspensions were subjected to repetitive cycles of freezing in liquid nitrogen and thawing in order to destroy multilamellar lipid layers. Liposomes were then formed at 55 °C by an extrusion process (46, 47) by using a mini-extruder system (Avanti Polar Lipids) and Teflon membranes (pore size:100 nm; Whatman). Electron microscopy showed that the resulting liposomes had a narrow size distribution with a diameter around 100 nm.

Coupling Reactions and Purification of Coupled PrP-SH (MS/HPLC Analysis)—Coupling reactions were performed in a volume of 2 ml at room temperature in 5 mM Tris/HCl, pH 8.0, for 12 h. Reactions for PC/Chol liposomes contained a total of 2 mM lipid and 10 µM PrP-SH, i.e. equimolar amounts of PrP-SH and PDP-DHPE coupling reagent available on the outer liposome surface. Because of a higher tendency of PrP-SH to aggregate in the presence of raft liposomes, the reaction volume was increased 2-fold (1 mM lipid and 5 µM PrP-SH) for synthesis of raft liposome-coupled PrP. Nevertheless, a fraction of PrP-SH aggregated in all coupling reactions. Aggregated material was removed by centrifugation at 25 °C and 14,000 x g for 30 min. Uncoupled PrP-SH was then separated from liposome-bound PrP-SS-PDHPE in the supernatant by an ultracentrifugation step in a Beckman OptimaTM MAX ultracentrifuge (TLA-55, 50,000 rpm, 4 °C, 20 min). Under these conditions, liposomes harboring PrP-SS-PDHPE molecules were pelleted, whereas uncoupled PrP-SH stayed in the supernatant. Liposomes were suspended in water. In the case of the raft liposomes, ultracentrifugation and subsequent suspension in water had to be repeated twice in order to completely remove uncoupled PrP-SH. Covalent coupling was verified by analytical reversed-phase HPLC and MALDI-TOF. HPLC separation was performed on a Vydac C18 column (4.6 x 250 mm) at 55 °C, where the uncoupled and coupled forms of PrP-SH gave well distinguishable peaks when an acetonitrile gradient from 0 to 95% (v/v) in 0.12% (v/v) trifluoroacetic acid was applied (flow rate, 0.8 ml/min). MALDI-TOF confirmed generation of PrP-SS-PDHPE (calculated mass, 24,417 Da; measured, 24,426 Da) from PrP-SH (calculated, 23,551 Da; measured, 23,556 Da).

Determination of Lipid/Protein Ratios—Lipid concentrations in liposomes containing PrP-SS-PDHPE were determined by two independent methods. The first assay was based on complex formation of phospholipids with ammonium ferrothiocyanate in organic solvents (48). Samples (50 µl) were mixed with methanol (100 µl), and 1 ml of chloroform and 0.8 ml of an aqueous ammonium ferrothiocyanate solution (27.03 g/liter FeCl3·6H2O, 30.4 g/liter NH4SCN) were added. After a vortexing step, the samples were centrifuged for 5 min at 5000 x g, and the absorbance of the chloroform phase was determined at 485 nm. Lipid concentrations were calculated from calibration curves for pure PC/Chol and raft liposomes, respectively. The second method exploited the linear correlation between liposome concentration and light scattering at 350 nm (A350) in the concentration range of 0.3–10 mM lipid. Calibration curves were determined for PC/Chol and raft liposomes, from which the lipid content of the coupled samples could be calculated. Results from both methods coincided completely.

The concentration of PrP-SS-PDHPE in liposomes was also determined by two independent assays. The first method was the BCA staining method performed with the BCA Promega kit. Calibration curves were recorded with wild type PrP-(23–231) in the presence of liposomes with known concentration to account for the influence of lipid on the absorbance at 562 nm. The concentration of PrP-SS-PDHPE was also measured by absorbance, using the extinction coefficient of uncoupled PrP-SH and correction for light scattering through liposomes.

CD Measurements—Far-UV CD spectra were measured on a Jasco 710 CD spectropolarimeter at 25 °C. All samples were centrifuged (15 min, 14,000 x g) prior to the measurements. Spectra were recorded in quartz cuvettes (0.1 or 0.02 cm when lipid concentrations exceeded 2.0 mM), accumulated 15 times, and corrected for the respective liposome/buffer solutions. Buffers used were 5 mM NaH2PO4/NaOH, pH 7.0, and 5 mM acetic acid/NaOH, pH 5.0. Because of the high aggregation tendency of raft liposome-coupled PrP-SS-PDHPE after addition of buffer and the comparably low protein concentrations, far-UV CD spectra of raft liposome-coupled PrP-SS-PDHPE were accumulated 75 times in 2 mM acetic acid/NaOH, pH 5.0. Protein concentrations for the far-UV CD spectra of PrP-(23–231) in the presence or of PrP-SS-PDHPE coupled to PC/Chol liposomes were between 6.5 and 8.8 µM, and lipid concentrations varied between 1.1 and 3.0 mM. Far-UV CD spectra of PrP-(23–231) in the presence or of PrP-SS-PDHPE coupled to raft liposomes were recorded at protein concentrations between 1.3 and 8.2 µM and lipid concentrations between 0.58 and 0.81 mM.

Limited Proteolysis with Trypsin and Proteinase K—Limited proteolysis was performed at 37 °C for 1 h at the following proteinase concentrations: 0, 0.0005, 0.005, 0.05, 0.5, 5, 50, 500 µg/ml proteinase K or 0, 0.0005, 0.005, 0.05, 0.5, 5, 50, 500, 1250 µg/ml trypsin. Samples testing the effect of PC/Chol liposomes on the stability of PrP-(23–231) and covalently coupled PrP-SS-PDHPE and the respective controls contained 7.2 µM protein and lipid concentrations of 2.5 mM in the case of coupled PrP-SS-PDHPE and 2.2 mM in the case of the uncoupled PrP-(23–231) in 20 µl of buffer (for trypsin digestion, 50 mM NH4HCO3/HCl, pH 7.8; for proteinase K digestion, 100 mM MOPS/NaOH, pH 7.4, 150 mM NaCl). Because of the aggregation tendency of samples containing raft liposomes, proteolysis was performed in 10-fold diluted, 200-µl samples containing 0.5 µM protein and 0.22 mM lipid in the case of coupled PrP-SS-PDHPE and uncoupled PrP-(23–231) in the presence of raft liposomes. After 1 h of incubation, reactions were quenched with phenylmethylsulfonyl fluoride, concentrated, and analyzed by SDS-PAGE. After blotting onto polyvinylidene difluoride membranes, the major proteolytic fragments were identified by Edman sequencing.

Electron Microscopy Analysis—Samples of liposomes in the presence and absence of PrP-(23–231) and with coupled PrP-SS-PDHPE were adsorbed to glow-discharged carbon-coated copper grids. These were washed twice with deionized water, negatively stained with 2% (w/v) uranyl acetate, and air-dried after removal of excess liquid. Specimens were examined in a Philips CM12 transmission electron microscope at 100 kV, and images were recorded with a Gatan 694 slow scan CCD camera.

Light Scattering Analysis—Samples of liposomes (10 µl, lipid concentration, 1 mM) were measured for 15 min at 25 °C in 0.15-cm cuvettes using a Protein Solutions DynaPro Molecular Sizing Instrument containing a Temperature Controlled Micro Sampler unit, in order to investigate liposomal integrity under various conditions (0–9 M urea; 0–5 M guanidinium chloride; after heating to 90 °C).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Covalent Coupling of PrP-SH to Liposomes—Initial attempts to refold and purify expressed PrP-SH from inclusion bodies isolated from the E. coli cytoplasm, as described previously for recombinant wild type PrP (40), were hampered by formation of disulfide-linked oligomers during oxidative refolding. The problem of nonspecific formation of intermolecular disulfides during refolding could be circumvented by construction of an N-terminally His6-tagged version of PrP-SH and oxidative refolding while bound to a Ni-NTA column. This procedure prevented formation of higher disulfide-linked oligomers and yielded a main fraction of monomeric correctly disulfide-bridged PrP-SH after refolding. After cleavage of the N-terminal His6 tag with thrombin, a small fraction of disulfide-linked PrP-SH dimers was removed with an additional ion exchange chromatography step (Fig. 2).



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  		FIG. 2.
Purification of PrP-SH. A non-reducing, Coomassie-stained SDS-gel is shown. Lane 1, inclusion body preparation; lane 2, elution from the Ni-NTA column; lane 3, pure PrP-SH after cleavage of the His6 tag; lane 4, concentrated sample of pure PrP-SH.

 
Purified PrP-SH was covalently coupled at pH 8.0 to either PC/Chol or raft liposomes with an average diameter of 100 nm, containing 1 mol % of the thiol-reactive phospholipid PDP-DHPE. PrP-SH was used in stoichiometric amounts relative to PDP-DHPE during the coupling reaction, assuming that only 50% of PDP-DHPE were available for coupling to liposomes (corresponding to an equal distribution of PDP-DHPE between the inner and outer leaflet of the membrane bilayer). The structure of the resulting PrP derivative linked to the phosphatidylethanolamine unit is shown in Fig. 1 and was termed PrP-SS-PDHPE. Coupling efficiency strongly decreased with increasing ionic strength in the coupling buffer because of irreversible, nonspecific aggregation of PrP. Best coupling yields were obtained at low ionic strength (5 mM Tris/HCl, pH 8.0). As PrP-SS-PDHPE could not be discriminated from PrP-SH by non-reducing SDS-PAGE (not shown), synthesis of the coupling product was verified by analytical reversed-phase HPLC and MALDI-TOF mass spectrometry (Fig. 3 and Fig. 4).



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  		FIG. 3.
HPLC profiles documenting the synthesis and purification of PrP-SS-PDHPE. a, purification of PrP-SS-PDHPE after coupling to PC/Chol liposomes; b, purification of PrP-SS-PDHPE after coupling to raft liposomes. In each panel the peak on the left (1) corresponds to uncoupled PrP-SH, and the peak on the right (2) represents PrP-SS-PDHPE. The several runs indicated by i–iv are as follows: (i) uncoupled PrP-SH alone, in the absence of liposomes; (ii) the total reaction mixture after 12 h of incubation; (iii) the supernatant after removal of aggregated material by low speed centrifugation; and (iv) resuspended and washed liposomes harboring PrP-SS-PDHPE after ultracentrifugation.

 



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  		FIG. 4.
MALDI-TOF spectra of PrP-SH alone (a) and purified PrP-SS-PDHPE coupled to PC/Chol liposomes (b). Calculated masses are indicated in brackets. Sinapinic acid was used as MALDI matrix.

 
The amount of coupling reagent used for all experiments described below was kept at 1 mol % PDP-DHPE to guarantee liposomal integrity and to avoid undesirable effects, e.g. increased aggregation tendencies, that have been reported previously for higher fractions of PDP-DHPE and similar coupling reagents (49, 50).

Purification of Liposomes Bearing Covalently Coupled PrP-SS-PDHPE—Even under conditions where nonspecific aggregation during coupling of PrP-SH was minimal, we could not achieve quantitative coupling to PDP-DHPE. The coupling reaction thus contained a mixture of liposome-coupled PrP-SS-PDHPE and uncoupled PrP-SH. Application of samples from the total reaction mixture to an HPLC column revealed that about one-third of PrP-SH did not react with PDP-DHPE and remained uncoupled in the case of PC/Chol liposomes; in the case of raft liposomes, where we observed lower coupling efficiency, about half of the PrP-SH molecules remained uncoupled (Fig. 3). Therefore, a procedure for separation of liposomes bearing covalently coupled PrP-SS-PDHPE from uncoupled PrP-SH had to be developed. We first removed nonspecific aggregates by a low speed centrifugation step at 14,000 x g for 30 min. HPLC analysis of the supernatant revealed that the amount of PrP-SS-PDHPE was decreased in the supernatant, independent of the liposome species involved, whereas the peaks corresponding to free PrP-SH remained unchanged (Fig. 3). The fraction of PrP-SS-PDHPE covalently coupled to liposomes could then be separated from uncoupled protein by an ultracentrifugation step in which the liposomes were sedimented. Washed liposome pellets were then suspended in water, and HPLC analysis showed that liposome-associated PrP-SS-PDHPE was pure and free of PrP-SH (Fig. 3).

By assuming that 50% of PDP-DHPE is incorporated into the outer liposome leaflet, and an average area of about 78.5 Å2 per head group of PC/Chol dimer (51), 400 surface-accessible PDP-DHPE molecules per liposome with an average diameter of 100 nm are expected. Analysis of the actual PrP/lipid ratios in purified PrP-SS-PDHPE-liposomes revealed, however, lower loading of the liposomes with PrP because of the incomplete coupling reaction described above. For PC/Chol preparations, the average molar PrP/total lipid ratio was around 0.0029 (instead of 0.005 for 100% coupling yield), corresponding to about 230 PrP molecules per liposome. Assuming an average diameter of 2.9 nm for the folded C-terminal PrP domain (35), this corresponds to a covering of about 5% of the liposome surface with the C-terminal PrP domain PrP-(121–231). The total surface area covered is, however, significantly larger for liposome-associated full-length PrP, because of the large exposed surface area of the unstructured N-terminal PrP segment 23–120 (36). The average PrP/lipid ratio determined for raft preparations was slightly lower, with a PrP/lipid ratio of 0.0024, corresponding to about 190 PrP molecules per liposome. To test whether liposomes stayed intact after coupling of PrP-SH and purification by ultracentrifugation, electron micrographs of freshly prepared PC/Chol and raft liposomes were compared with those of purified liposomes bearing covalently bound PrP-SS-PDHPE. The overall appearance of non-loaded liposomes was indistinguishable from that of PrP-SS-PDHPE-loaded liposomes (Fig. 5). Moreover, no nonspecific aggregates were present in purified PrP-SS-PDHPE-loaded liposome solutions. As a control, it was also shown that addition of soluble wild type PrP-(23–231) to liposome preparations had no influence on the shape of PC/Chol and raft liposomes (Fig. 5).



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  		FIG. 5.
Electron micrographs of PC/Chol liposomes (a), PC/Chol liposomes in the presence of soluble PrP-(23–231) (b), PC/Chol liposomes with coupled PrP-SS-PDHPE (c), raft liposomes (d), raft liposomes in the presence of soluble PrP-(23–231) (e), and raft liposomes with coupled PrP-SS-PDHPE (f).

 
Characterization of Liposome-bound PrP by Far-UV CD Spectroscopy—The influence of the covalent coupling to membranes on the conformation of the full-length prion protein was first investigated by far-UV CD spectroscopy. The spectra for full-length PrP-(23–231) in solution were compared with those of soluble, full-length PrP-(23–231) in the presence of liposomes and liposome-associated PrP-SS-PDHPE. Soluble PrP-SH exhibits a far-UV CD spectrum identical to that of wild type PrP-(23–231) (data not shown).

Spectra were recorded at pH 7.0 and pH 5.0, i.e. roughly the pH range encountered by PrP when present on the cell surface or after incorporation into endosomes, and corrected for the respective liposome/buffer solutions. The PC/Chol liposomes, consisting of zwitterionic phosphatidylcholine and cholesterol only and therefore carrying no net charge, neither had an influence on the far-UV CD spectra of uncoupled full-length PrP-(23–231) nor on the far-UV CD spectra of liposome-bound PrP-SS-PDHPE. At both pH 7.0 and pH 5.0, the far-UV CD spectra of PrP-(23–231), of PrP-(23–231) in the presence of PC/Chol liposomes, and of liposome-bound PrP-SS-PDHPE coincide completely and show the typical minima at 208 and 222 nm, indicative of the typical {alpha}-helical structure of PrPC (Fig. 6). Spectroscopic analysis of raft-coupled PrP-SS-PDHPE was hampered by a relatively high tendency of the preparation to aggregate at concentrations required for CD analysis. Although spectra at pH 5.0 and pH 7.0 could be recorded for a mixture of soluble PrP-(23–231) and raft liposomes, the lipid concentrations had to be 3–4-fold lower compared with the solutions containing PC/Chol liposomes to prevent nonspecific aggregation. Recording of far-UV CD spectra of raft-coupled PrP-SS-PDHPE at neutral pH was prevented by complete aggregation. At pH 5.0, far-UV CD spectra of coupled PrP-SS-PDHPE could, however, be recorded at very low ionic strength (2 mM acetic acid/NaOH). All spectra showed that the overall {alpha}-helical structure of PrP was also retained in the presence of raft liposomes. However, covalent coupling of PrP to raft liposomes caused a slight spectral shift toward more negative molar mean residue ellipticities. In contrast, uncoupled PrP-(23–231) in the presence of raft liposomes exhibited far-UV CD spectra identical to PrP-(23–231) in the absence of liposomes (Fig. 6).



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  		FIG. 6.
Far-UV CD spectra of PrP-(23–231) (crosses), soluble PrP-(23–231) in the presence of PC/Chol liposomes (2.20 mg lipid/ml; empty circles), PrP-SS-PDHPE coupled to PC/Chol liposomes (2.27 mg lipid/ml; empty diamonds) at pH 5.0 (a) and pH 7.0 (b), and far-UV CD spectra of PrP-(23–231) (crosses) and soluble PrP-(23–231) in the presence of raft liposomes (0.55 mg lipid/ml; empty circles) and PrP-SS-PDHPE coupled to raft liposomes (0.58 mg lipid/ml; empty diamonds) at pH 5.0 (c).

 
Stability of Membrane-bound PrP—Light scattering experiments revealed that liposomes were not stable at denaturant concentrations above 4 M urea and above 2.5 M GdmCl. The influence of a lipid membrane environment on the stability of full-length PrP could therefore not be assessed by urea or GdmCl-induced folding transitions, as the full-length prion protein is not completely unfolded at denaturant concentrations at which the liposomal integrity is no longer given (43, 52). The same holds true for thermal unfolding transitions, as the liposomes became unstable at temperatures above 75 °C, and the full-length prion protein is not completely unfolded at this temperature (29).

The stability of PrP-(23–231) in the absence and presence of liposomes, and of liposome-coupled PrP-SS-PDHPE, was then investigated by resistance against proteolysis by proteases with different specificities. This method makes use of the fact that protein stability correlates with protease resistance, as the unfolded state, which is populated to a higher fraction in a less stable protein, is preferably degraded by proteases and removed from the folding equilibrium (53). Limited proteolysis was performed for 1 h at 37 °C in the presence of increasing concentrations of either proteinase K or trypsin. No differences in the digestion pattern were observed for recombinant PrP-(23–231), recombinant PrP-(23–231) in the presence of PC/Chol, or raft liposomes and PrP-SS-PDHPE (Fig. 7). This indicates that neither the presence of PC/Chol or raft liposomes nor the covalent coupling of PrP to PC/Chol or raft liposomes destabilized PrP-SS-PDHPE relative to soluble, wild type PrP-(23–231).



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  		FIG. 7.
Non-reducing, Coomassie-stained SDS-gels of limited proteolysis assays, with trypsin (a) and with proteinase K (b). Increasing concentrations of protease are indicated by the black block wedges above the gels. Specific concentrations are as follows: lane 1, 0 µg/ml; lane 2, 0.0005 µg/ml; lane 3, 0.005 µg/ml; lane 4, 0.05 µg/ml; lane 5, 0.5 µg/ml, lane 6, 5 µg/ml; lane 7, 50 µg/ml; lane 8, 500 µg/ml; and lane 9, 1250 µg/ml. S, molecular mass standard. Bands marked by arrows in the corresponding upper left panels were identified by N-terminal Edman sequencing (see Table I).

 
In accordance with the known three-dimensional structure of full-length murine PrP-(23–231) (37), Edman sequencing of the major proteolysis products of PrP-(23–231) and PrP-SS-PDHPE yielded the folded C-terminal domain of PrP, PrP-(121–231), and the N-terminally truncated fragments of the unstructured N-terminal tail segment 23–120 for both digestion with proteinase K and digestion with trypsin (Table I). This demonstrates that the segment 23–120 is not protected against proteolysis by the membrane environment in liposome-bound PrP-SS-PDHPE, and that the stability and structure of the C-terminal domain 121–231 of PrP-SS-PDHPE is comparable with that in the context of free PrP-(23–231). The slightly different far-UV CD spectrum observed for PrP-SS-PDHPE coupled to raft liposomes (Fig. 6c) is therefore most likely not indicative of a significant conformational change.


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  		TABLE I
Protease cleavage sites determined by Edman sequencing of the degradation products indicated in Fig. 8

Sequencing results obtained for degradation products of PrP-(23–231) in the absence and presence of liposomes or in samples with liposome-bound PrP-SS-PDHPE were identical.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
One of the key events in prion pathogenesis, the conversion from the benign cellular prion protein, PrPC, to its scrapie isoform, PrPSc, involves a structural transition from a mainly {alpha}-helical conformation to a conformation with an increased portion of {beta}-sheet structure (6–11). In order to elucidate the molecular mechanism underlying this conversion reaction, thermodynamic and structural studies on the prion protein have been performed with soluble, recombinant PrP from E. coli that is assumed, due to coincident biophysical characteristics, to correspond to the structure of the PrPC isoform of the prion protein (7, 40, 52).

In the study presented here, we focused on the fact that PrPC is covalently attached to cellular membranes in vivo via a C-terminal GPI anchor (16), and that the conversion of PrPC to PrPSc most likely occurs on the membrane surface. An experimental model was developed that allows for the first time the monitoring of the impact of covalent membrane attachment on the structure and stability of recombinant prion protein. The results for recombinant full-length prion protein covalently coupled to membranes clearly indicate that covalent membrane attachment via a relatively flexible module like the GPI anchor (mimicked in this study by a glycine linker) does not lead to changes in the overall structure and the overall stability of the protein. Although for full-length PrP coupled to raft-like membranes a slight increase was observed in the amplitude of the far-UV CD signal upon covalent attachment of PrP to raft liposomes, this change in signal was not accompanied by a partial protection of the membrane-coupled protein against digestion with proteases. These spectroscopic data, together with the complete agreement of the proteolytic pattern of membrane-coupled PrP-SS-PDHPE and soluble PrP-(23–231), suggest that raft lipids do not change the structure and stability of the cellular prion protein, indicating that the structures determined by NMR for the soluble recombinant prion proteins of mouse, hamster, man, and cattle (32–39) might well represent the conformation adopted by PrPC on the cell surface in the absence of ligands like copper ions (54, 55) or other interaction partners (56). An alternative explanation for the requirement of raft localization of the prion protein for its conversion from PrPC to PrPSc, as observed previously in cell culture models (17, 23, 24, 26, 27), may therefore be an increase in local, effective PrPC concentration on the outer surface of the membrane that facilitates PrP-PrP interactions. Another possible mechanism might involve interactions of PrPC with other raft-localized factors, for example the postulated host factor protein X (56).

Our present investigation was prompted by previous studies on soluble, recombinant prion proteins without lipid anchor in the presence of lipid membranes (29–31). Morillas et al. (29) could demonstrate that binding of PrP to lipid membranes required negatively charged lipids and was accompanied by the formation of structure in the previously unstructured N-terminal tail of the full-length prion protein. Furthermore, the C-terminal domain was found to be destabilized in the presence of acidic membranes, especially at acidic pH values (29). In contrast, incubation of the prion protein with vesicles consisting solely of zwitterionic phosphatidylcholine did not result in observable binding of recombinant PrP to the respective membranes. In another study by Sanghera and Pinheiro (30), it was additionally demonstrated that PrP could associate with uncharged membranes in the gel-phase state consisting exclusively of dipalmitoylphosphatidylcholine or a raft-like lipid composition under neutral pH conditions. Both studies, however, could not exclude that the prion protein associated abnormally with the model membranes, whereas a natural orientation relative to the membrane surface was guaranteed in our present study. We also considered the size and flexibility of the natural GPI anchor of the prion protein in the design of our synthetic membrane anchor (57–60), which differs from the natural GPI anchor only by replacement of the central core oligosaccharide by a flexible linker peptide of six amino acids. The length of this linker peptide was adjusted to roughly equal the average distance of 20 Å between the C-terminal carboxylate of PrP (Ser-231) and the phosphate of the phosphatidyl group (three-dimensional reconstruction of the minimal core structure of GPI anchors, D-Man{alpha}(1–2)-D-Man{alpha}(1–6)-D-Man{alpha}(1–4)-D-GlcNAc-(1–6)-inositol, at www.dkfz-heidelberg.de/spec/sweet2). We thus expect the same distance from the membrane and orientation relative to the membrane as in natural PrPC (60). Besides these structural considerations, the fact that the outer leaflets of biological membranes are usually devoid of negatively charged phospholipids was accounted for by exclusive use of uncharged lipid membranes in the liquid-ordered phase, i.e. the state that raft microdomains are supposed to adopt in cellular membranes (20, 21, 44). In accordance with the results for soluble PrP in the presence of model membranes (29, 30), we found that the {alpha}-helical character of full-length PrP was retained upon covalent membrane attachment to uncharged liposomes. However, we did not observe an increase in {alpha}-helical structure for soluble PrP in the presence of PC/Chol or raft liposomes, as was reported previously for PrP-(90–231) (30). These different results may be due to the N-terminally truncated PrP constructs used in previous studies (i.e. PrP-(90–231) versus PrP-(23–231) used in the present study) or due to differences in the respective membrane model systems. A slightly more negative CD signal was only observed for PrP covalently bound to raft liposomes. We do not consider this difference an indication for structure formation in segment 23–120 of membrane-associated PrP, not only because recording of CD spectra of raft liposome-bound PrP was hampered by protein aggregation at the concentrations that had to be used for CD spectroscopy (see above), but particularly because limited proteolysis yielded the same high protease sensitivity for segment 23–120 in membrane-bound PrP compared with free PrP. Furthermore, we have no indication for a disruption of the bilayer by membrane-bound PrP from the respective electron micrographs that would support the idea of a PrP insertion into the membrane, as observed previously for binding of PrP to negatively charged membranes (29–31, 61).

The role of membrane attachment for the transition from PrPC to protease-resistant PrP was also investigated previously (27, 28) by in vitro conversion experiments. Protease-sensitive PrP lacking the GPI anchor was co-fractionated with raft-like membranes in a binding assay using floatation on density gradients. In agreement with our conclusion that the neighborhood of raft-like membranes does not significantly influence the structure and stability of PrPC, this mode of GPI anchor-independent, non-covalent membrane attachment allowed the conversion to protease-resistant PrP by external PrPSc with almost identical conversion efficiencies as observed for PrP without GPI anchor in the absence of membranes (28). The fact that we did not detect binding of soluble, recombinant PrP to raft-like liposomes in our present study might be due to the significantly higher ionic strengths used in the experiments by Baron and Caughey (28). Whether the GPI anchor-independent mode of membrane association of PrPC observed by Baron and Caughey (28) is relevant for the conversion from PrPC to PrPSc in vivo remains an open question, as the majority of PrPSc isolated from the brains of infected Syrian hamsters contains a GPI anchor (16, 58).

Most interesting, protease-sensitive PrPC with a GPI anchor could not be converted to the protease-resistant form by exogenous PrPSc in in vitro conversion experiments unless the GPI anchor was either cleaved off or PrPSc was incorporated into the same membrane by use of a membrane-fusing agent (28). This suggests that co-localization of PrPC and PrPSc in contiguous membranes is a critical prerequisite for prion propagation in vivo. In this context, our model system guarantees that at least 200 molecules of protease-sensitive PrP would be localized in the same membrane as PrPSc after insertion of a single PrPSc oligomer into a synthetic PrP-SS-PDHPE liposome. This provides a good basis for systematic future in vitro conversion experiments with covalently membrane-coupled, recombinant PrP to further investigate the molecular basis of PrPSc formation.


   FOOTNOTES
 
* The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

To whom correspondence should be addressed: Institut für Molekularbiologie und Biophysik, Eidgenössische Technische Hochschule Hönggerberg, CH-8093 Zürich, Switzerland. Tel.: 41-1-6336819; Fax: 41-1-6331036; E-mail: rudi{at}mol.biol.ethz.ch.

1 The abbreviations used are: TSE, transmissible spongiform encephalopathy; Chol, cholesterol; GPI anchor, glycosylphosphatidylinositol anchor; MALDI-TOF, matrix-assisted laser desorption time-of-flight mass spectrometry; PC, phosphatidylcholine; PDP-DHPE, N-((2-pyridyldithio)-propionyl)-1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine, triethylammonium salt; PrPC, cellular isoform of the prion protein; PrPSc, scrapie isoform of the prion protein; PrP-SH, recombinant, murine full-length PrP with a C-terminal Gly5Cys-linker; PrP-SS-PDHPE, PrP-SH after covalent coupling to PDP-DHPE; Ni-NTA, nickel-nitrilotriacetic acid; GdmCl, guanidinium chloride; HPLC, high pressure liquid chromatography; MOPS, 4-morpholinepropanesulfonic acid. Back



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HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
 All ASBMB Journals   Molecular and Cellular Proteomics 
 Journal of Lipid Research   ASBMB Today 
Copyright © 2004 by the American Society for Biochemistry and Molecular Biology.
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