©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
Sulfated Glycans Stimulate Endocytosis of the Cellular Isoform of the Prion Protein, PrP, in Cultured Cells (*)

(Received for publication, August 7, 1995; and in revised form, October 5, 1995)

Show-Ling Shyng (§) Sylvain Lehmann (¶) Krista L. Moulder David A. Harris (**)

From the Department of Cell Biology and Physiology, Washington University School of Medicine, St. Louis, Missouri 63110

ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

There is currently no effective therapy for human prion diseases. However, several polyanionic glycans, including pentosan sulfate and dextran sulfate, prolong the incubation time of scrapie in rodents, and inhibit the production of the scrapie isoform of the prion protein (PrP), the major component of infectious prions, in cultured neuroblastoma cells. We report here that pentosan sulfate and related compounds rapidly and dramatically reduce the amount of PrP^C, the non-infectious precursor of PrP, present on the cell surface. This effect results primarily from the ability of these agents to stimulate endocytosis of PrP^C, thereby causing a redistribution of the protein from the plasma membrane to the cell interior. Pentosan sulfate also causes a change in the ultrastructural localization of PrP^C, such that a portion of the protein molecules are shifted into late endosomes and/or lysosomes. In addition, we demonstrate, using PrP-containing bacterial fusion proteins, that cultured cells express saturable and specific surface binding sites for PrP, many of which are glycosaminoglycan molecules. Our results raise the possibility that sulfated glycans inhibit prion production by altering the cellular localization of PrP^C precursor, and they indicate that endogenous proteoglycans are likely to play an important role in the cellular metabolism of both PrP^C and PrP.


INTRODUCTION

Prion diseases are a group of transmissible, neurodegenerative disorders including Creutzfeldt-Jakob disease, kuru, Gerstmann-Sträussler syndrome, and fatal familial insomnia in human beings, as well as bovine spongiform encephalopathy and scrapie in animals (reviewed by Gajdusek(1990) and Prusiner and DeArmond (1994)). The major component of infectious prion particles is a protein called PrP(^1)(Prusiner et al., 1984; Bolton et al., 1987). PrP is a posttranslationally modified isoform of PrP^C, a glycolipid-anchored, plasma membrane protein of unknown function that is widely expressed on neurons and glia in the central nervous system (Oesch et al., 1985; Stahl et al., 1987; Harris et al., 1993a). Prion replication is thought to involve conversion of endogenous PrP^C into infectious PrP, although the cellular and molecular details of this process are poorly understood (Prusiner et al., 1990; Büeler et al., 1993; Kocisko et al., 1994). Recent evidence suggests that the two isoforms differ in their three-dimensional conformation, and that transformation of alpha-helices into beta-sheets plays a role in formation of PrP (Caughey et al., 1991a; Safar et al., 1993; Pan et al., 1993).

There are currently no effective therapies for human prion diseases, although several chemotherapeutic agents have been tested in animal models (Pocchiari et al., 1991; Ingrosso et al., 1995). Polyanionic glycans such as pentosan sulfate (PS) and dextran sulfate have been among the most intensively studied (Ehlers and Diringer, 1984; Kimberlin and Walker, 1986; Farquhar and Dickinson, 1986; Diringer and Ehlers, 1991; Ladogana et al., 1992). These agents were initially tested because they were known to be active against conventional DNA and RNA viruses, but it was found that they were also effective in vivo against infection by scrapie prions, prolonging the incubation time, and in some cases, completely preventing the development of symptoms when administered prophylactically to mice and hamsters.

Caughey and colleagues have recently investigated the mechanism of this effect at the cellular level. They showed that the disulfonated, amyloid-binding dye Congo red, as well as glycans such as pentosan sulfate and dextran sulfate inhibit prion replication and PrP accumulation in scrapie-infected neuroblastoma cells (Caughey and Race, 1992; Caughey et al., 1993; Caughey and Raymond, 1993); similar results have also been obtained by Gabizon et al.(1993) using low molecular weight heparin. These effects were proposed to result from a direct interaction between the inhibitors and PrP, based on the observation that PrP^C binds in vitro to beads containing immobilized Congo red or heparin (Caughey et al., 1994). Because the agents tested are structurally related to glycosaminoglycans (GAGs) synthesized by cells, these workers have further suggested that PrP^C and/or PrP bind to endogenous GAG chains, and that competitive inhibition of this association is responsible for the negative effect of the compounds on prion synthesis (Priola and Caughey, 1994). The proposal that PrP interacts with cellular GAGs is consistent with the fact that amyloid plaques in scrapie-infected brain often contain sulfated proteoglycans (Snow et al., 1990; Gulroy et al., 1991). However, direct evidence for the existence of GAG-related binding sites on cells for either PrP^C or PrP has been lacking.

Also unknown is how sulfated glycans and Congo red affect the cellular trafficking of PrP. This is an important question, since alterations in the trafficking and localization of either PrP^C or PrP might alter the efficiency of prion production. Recent studies of scrapie-infected neuroblastoma cells suggest that conversion of PrP^C to PrP takes place either on the plasma membrane or along an endocytic pathway (Caughey et al., 1991b; Caughey and Raymond, 1991; Borchelt et al., 1992; Taraboulos et al., 1992). Our own studies strongly support this conclusion, since we have shown that PrP^C constitutively cycles between the cell surface and an endocytic compartment, with a transit time of 60 min in cultured neuroblastoma cells (Shyng et al., 1993), and that internalization of PrP^C is mediated by clathrin-coated pits (Shyng et al., 1994). These results identify early endosomes and clathrin-coated vesicles as potential sites for the generation of PrP. Conceivably, chemotherapeutic agents could act by redistributing PrP^C or PrP away from these or other relevant cellular compartments.

To understand how sulfated glycans act to prevent prion production in cultured cells, we have examined the effect of these compounds on the cellular trafficking of PrP^C, the precursor of PrP, with particular emphasis on the endocytic pathway. We report here that PS, as well as several other polyanions, dramatically decrease the amount of PrP^C on the surface of neuroblastoma cells by enhancing its rate of endocytosis. PS also redistributes a portion of the PrP molecules from early to late endocytic compartments. In addition, we demonstrate, using PrP-containing bacterial fusion proteins, that cultured cells express GAG-containing surface binding sites for PrP. Our results raise the possibility that sulfated anions inhibit prion synthesis by altering the cellular localization of the PrP^C precursor, and they suggest that endogenous proteoglycans are likely to play an important role in the cellular metabolism of both PrP^C and PrP.


MATERIALS AND METHODS

Reagents and Antibodies

Rabbit antisera raised against bacterial fusion proteins encompassing amino acids 35-96 or 144-220 of chPrP (Harris et al., 1993b) were used to recognize wild-type and Delta25-91 chPrPs, respectively. A rabbit antiserum raised against a synthetic peptide encompassing amino acids 45-66 of mouse PrP was used to recognize mouse PrP (Shyng et al., 1995). A rat monoclonal antibody (Ig11) raised against the mouse lysosomal membrane protein lgp120 was obtained from Hugh Rosen (Merck). Cell culture reagents were from the Tissue Culture Support Center at Washington University. Phosphatidylinositol-specific phospholipase C (PIPLC) was prepared as described previously (Shyng et al., 1995). Amylose resin was obtained from New England Biolabs, and gold-conjugated secondary antibodies from Jackson Immunoresearch. Heparitinase (heparin lyase III) was purchased from Sigma or ICN, and N-glycosidase F from Boehringer Mannheim. All other reagents were from Sigma.

Cell Lines

This study employed untransfected mouse N2a neuroblastoma cells (ATCC CCL131), as well as stably transfected clones of N2a cells expressing either wild-type chPrP (line A26), or Delta25-91 chPrP (Harris et al., 1993b; Shyng et al., 1995). Cells were maintained in minimal essential medium supplemented with 10% fetal calf serum, non-essential amino acids, and penicillin/streptomycin in an atmosphere of 5% CO(2), 95% air. Chinese hamster ovary (CHO) cells deficient in xylose transferase (clone 745), as well as wild-type CHO K1 cells, were kindly provided by Dr. Jeffrey Esko (Esko, 1991).

Immunoblotting, Metabolic Labeling, and Immunoprecipitation

To release cell surface PrP, cultures were treated with PIPLC (1 unit/ml) in Opti-MEM at 4 °C for 2 h. Proteins were precipitated from the PIPLC incubation medium with at least four volumes of methanol at -20 °C, collected by centrifugation, and immunoblotted with anti-PrP antibodies using ECL for visualization. Films were digitized using an HP ScanJet II scanner, and band intensities quantitated using SigmaScan/Image (Jandel Scientific).

For metabolic labeling, confluent cultures were incubated for 30 min in serum-free Opti-MEM lacking methionine, and then for 30 min in the same medium containing ICN TranS-Label (250 µCi/ml, 1,000 Ci/mmol). In some cases, cells were chased in Opti-MEM (Life Technologies, Inc.), and were treated with PIPLC as described above. Cell lysates were incubated with 0.01 units/ml N-glycosidase F at 37 °C for 16 h to cleave N-linked oligosaccharides, and PrP immunoprecipitated as described previously (Harris et al., 1993b). Radiolabeled PrP was quantitated by imaging of SDS-PAGE gels using a PhosphorImager (Molecular Dynamics, Sunnyvale, CA).

PrP Internalization Assays

Endocytosis of PrP was quantitated using either surface biotinylation or surface iodination, both as described previously (Shyng et al., 1995).

I-Transferrin Uptake Assay

Cells were incubated for 1 h at 4 °C in PBS containing I-transferrin (2 µM), and after washing, they were transferred to Opti-MEM containing 100 nM unlabeled transferrin with or without PS for 30 min at 37 °C. Prior to lysis, cells were treated with 0.25% Pronase to remove surface I-transferrin. The amount of internalized I-transferrin was determined by counting the radioactivity in the cell lysates.

Immunogold Labeling and Electron Microscopy

A26 cells were labeled with antibodies against chPrP 35-96 for 2 h at 4 °C and were then warmed to 37 °C for 1 h in the presence or absence of PS. Cells were then washed with PBS, fixed in 2% paraformaldehyde, 0.2% glutaraldehyde for 2 h, and scraped off the dish using a rubber policeman. The suspended cells were then collected by centrifugation, washed in PBS, and pelleted through 10% gelatin. Cells were infused with 2.3 M sucrose for 48 h at 4 °C, and after freezing in liquid nitrogen, 100-nm cryosections were cut at -90 °C on a Reichert-Jung UltraCut equipped with an FC4E Cryosystem (Leica). Sections were incubated sequentially with goat anti-rabbit antibodies conjugated to 6-nm gold particles, a rat monoclonal antibody against lgp120, and goat anti-rat antibodies coupled to 12-nm gold particles.

For quantitating colocalization of PrP and lgp120, 6- and 12-nm gold particles in intracellular vesicles were counted in 40 cells; 700 particles of each kind were counted for each experimental treatment. Vesicles were grouped according to the number of 12-nm gold particles they contained, and the percentage of total PrP particles associated with each group was calculated.

Bacterial Fusion Proteins

DNA fragments encoding chPrP residues 25-241, 25-116, and 117-241 were amplified by PCR from a full-length chPrP cDNA. The 5` primers used for PCR included an EcoRI restriction site followed by nucleotides encoding a Factor Xa cleavage site (5`-GAATTCTAATCGAGGGAAGG-3`). The 3` primers included a HindIII restriction site followed by a stop codon (5`-AAGCTTTCA-3`). The amplified DNA fragments were cut with EcoRI and HindIII and cloned first into the vector pGEX-KG (Guan and Dixon, 1991). The resulting plasmids were then cut with BamHI and HindIII to isolate a fragment that included the original PCR product plus a glycine linker region that is part of the pGEX-KG vector. This fragment was then cloned into the vector pMAL-c2 (New England Biolabs), which fuses the protein of interest to the Escherichia coli maltose-binding protein.

Fusion proteins were purified from ultrasonically disrupted bacteria to 95% homogeneity, using an amylose resin column according to the manufacturer's directions. Sodium deoxycholate (1% final concentration) was added to the purified fusion proteins to promote their solubilization, and the detergent was then removed by overnight dialysis at 4 °C against 20 mM HEPES (pH 8). Fusion proteins were iodinated using lactoperoxidase to a specific activity of 1-10 Ci/mmol.

Binding Assay

Confluent N2a or CHO cells in 12- or 24-well plates were washed twice in cold PBS, once with PBS containing 3% bovine serum albumin, and incubated with iodinated fusion proteins for 2 h at 4 °C. Cells were then washed with PBS/bovine serum albumin, lysed in SDS buffer, and the amount of bound fusion protein determined by counting. Cell lysates were analyzed by SDS-PAGE to confirm that the bound fusion protein was intact. Unlabeled fusion protein was used in the experiment shown in Fig. 10B, and the amount bound to cells was determined by immunoblotting cell lysates with anti-chPrP antibody, followed by quantitation of PrP bands on ECL films as described above.


Figure 10: Binding of MBP-chPrP 25-241 to N2a cells is reduced by treatments that interfere with GAGs. Panel A, binding assays were performed with 20 nM iodinated fusion protein as described in Fig. 9. After incubation with the fusion protein, some cells were washed with PBS containing 2 M NaCl (bar labeled NaCl). Other cells were treated with heparitinase (5 mIU/ml) for 2 h at 37 °C prior to incubation with fusion protein (bar labeled HSase). Data are expressed as a percentage of binding under control conditions. Nonspecific binding, measured in the presence of 4 µM unlabeled fusion protein, was first subtracted. Each bar shows the mean ± S.D. of values from 4 cultures. Panel B, binding assays were performed using 20 nM unlabeled fusion protein in the presence of PS, heparin (H), heparin sulfate (HS), or chondroitin sulfates (CS) (mixture of A, B, and C forms), all at 10 µg/ml. The amount of bound fusion protein was quantitated by immunoblotting cell lysates with anti-chPrP antibody. Data are expressed as a percentage of binding in the absence of drug. Each bar shows the mean and range of values for duplicate cultures.




Figure 9: Binding of iodinated chPrP fusion proteins to N2a cells. Panel A, N2a cells were incubated for 2 h at 4 °C with the indicated concentrations of I-labeled MBP-chPrP 25-241, and after washing, the amount of bound radioactivity determined by counting (curve marked Total). The binding reaction was also carried out in the presence of 4 µM unlabeled MBP-chPrP 24-241 (curve marked Nonspecific). The curve marked Specific is the difference between the Total and the Nonspecific curves. Each data point represents the average of triplicate determinations. Inset, Scatchard plot of the specific binding data shown in the main panel, giving a calculated K of 230 nM, with 8.8 times 10^5 sites/cell. Panel B, binding assays were performed as in panel A, using MBP-chPrP 25-116 and MBP-chPrP 117-241. The curves show specific binding. Inset, Scatchard plot of the binding data for MBP-chPrP 25-116 shown in the main panel, giving a calculated K of 240 nM, with 2.8 times 10^5 sites/cell.




RESULTS

PS Reduces the Amount of PrP Present on the Cell Surface

We first used immunofluorescence staining to analyze A26 cells, which are a stably transfected line of N2a neuroblastoma cells that express chPrP, the chicken homologue of mammalian PrP^C (Harris et al., 1993b). Incubation of cells for 12 h with 100 µg/ml PS caused a dramatic reduction in the amount of cell surface staining for chPrP (Fig. 1). This reduction was confirmed biochemically by using immunoblotting to quantitate the amount of PrP released by incubation of intact cells with phosphatidylinositol-specific phospholipase C (PIPLC), an enzyme that releases glycosyl phosphatidylinositol-anchored molecules from the cell surface (Fig. 2A). Treatment of cells with as little as 1 µg/ml PS for 12 h produced a detectable decrease in the amount of PIPLC-releasable chPrP, and a concentration of 100 µg/ml reduced the amount of chPrP by over 90%. The effect of PS was not restricted to chPrP, since the drug also reduced the amount of endogenous mouse PrP that could be labeled by surface iodination of N2a cells (Fig. 2B).


Figure 1: PS reduces the amount of cell surface PrP detected by immunofluorescence staining. A26 cells were incubated for 12 h at 37 °C in Opti-MEM with (PS) or without (Cont.) 0.1 mg/ml pentosan sulfate. They were then rinsed in PBS, incubated with anti-chPrP antibodies at 4 °C for 2 h, and fixed for 5 min in methanol at -20 °C. Fluorescein isothiocyanate-conjugated secondary antibodies were then applied, and the cells were viewed by fluorescence microscopy. Scale bar = 30 µm.




Figure 2: PS reduces the amount of PrP released by incubation of N2a cells with PIPLC. Panel A, A26 cells were incubated in Opti-MEM containing the indicated concentrations of PS for 12 h. Cells were then treated with PIPLC for 2 h at 4 °C to cleave cell surface PrP. Proteins were methanol-precipitated from the PIPLC incubation medium, and immunoblotted using anti-chPrP antibodies. Panel B: Untransfected N2a cells were incubated in Opti-MEM in the presence or absence of 100 µg/ml PS for 12 h. Cells were then surface-iodinated at 4 °C using lactoperoxidase, and mouse PrP immunoprecipitated from cell lysates and analyzed by SDS-PAGE.



Fig. 3shows a dose-response curve for the effect of PS. The drug caused marked reduction of cell surface chPrP at concentrations of 10 µg/ml or greater. This effect was observable with as little as 1 h of treatment, although it was more pronounced after 24 or 48 h. When present in the culture medium for these longer periods, PS caused a small (15-20%) but reproducible decrease in the amount of surface PrP at concentrations as low as 1 ng/ml; a similar result was obtained after 4 days of treatment (not shown). The concentration range of PS tested does not have any obvious effect on the morphology or growth rate of the cells.


Figure 3: Dose response of the reduction in cell surface PrP caused by PS. A26 cells were incubated in the indicated concentrations of PS for 1, 24, or 48 h. Cells were then treated with PIPLC for 2 h at 4 °C to cleave cell surface PrP. Proteins were methanol-precipitated from the PIPLC incubation medium and immunoblotted using anti-chPrP antibodies. The amount of chPrP was quantitated on digitized images of the ECL films and was expressed as percentage of the amount from control cells that were not treated with PS. Each data point represents average of duplicate samples.



We found that other polyanions also reduced the amount of chPrP on the surface of N2a cells (Fig. 4). Congo red and dextran sulfate of 500 kDa were about as effective as PS, dextran sulfate of 5 kDa was less potent, while chondroitin sulfate C was without effect. In additional experiments, heparin was about as potent as a 5-kDa dextran sulfate, and chondroitin sulfates A, B, and C were all ineffective at concentrations as high as 1 mg/ml (data not shown).


Figure 4: Other polyanionic compounds reduce the amount of cell surface PrP. A26 cells were untreated (Control), or were incubated for 24 h with PS (PS), Congo red (CR), dextran sulfate (DS) of 500 or 5 kDa, or chondroitin sulfate C (CSC), each at a concentration of 10 µg/ml. Cells were then treated for 2 h with PIPLC at 4 °C, and proteins in the PIPLC incubation medium were methanol-precipitated and immunoblotted with anti-chPrP antibodies.



PS Has No Effect on the Synthesis or Degradation of PrP

Several mechanisms could account for the effect of PS on surface PrP in cultured N2a cells, including changes in synthesis or degradation of the protein, or alterations in its distribution between the inside and outside of the cell. Another potential mechanism, increased release of surface PrP into the extracellular medium, is unlikely to be involved, since we found that PS actually decreases the amount of PrP in the medium (data not shown).

To assess an effect on synthetic rate, we measured the amount of chPrP that was metabolically labeled during a 30 min pulse with [S]methionine in the presence or absence of 1 mg/ml PS. The 30 min labeling period was chosen because at this concentration of PS, an effect on cell surface PrP is seen as early as 15 min (data not shown). Fig. 5A shows that the amount of PrP synthesized in the presence or absence of PS is not significantly different. PS also has no significant effect on the kinetics of transport of PrP to the cell surface during a chase period (data not shown).


Figure 5: PS does not affect the synthesis or degradation of PrP. Panel A, A26 cells were preincubated in medium without methionine for 30 min, and were then labeled with TranS-Label for 30 min in the presence of the indicated concentrations of PS. chPrP was immunoprecipitated from cell lysates, run on SDS-PAGE, and quantitated using a PhosphorImager. The amount of radiolabeled chPrP from PS-treated cells was expressed as a percentage of the amount from untreated cells. Each bar shows the mean and range of values for duplicate cultures. Panel B, A26 cells were labeled for 30 min with TranS-Label in the absence of PS, and chased in methionine-free medium in the absence or presence of PS (1 mg/ml) for 0, 2, 4, or 8 h. chPrP was immunoprecipitated from cell lysates, run on SDS-PAGE, and quantitated using a PhosphorImager. The amount of radiolabeled chPrP at each time point was expressed as a percentage of the amount present at the end of the labeling period (0 h of chase). Each point represents the average of duplicate samples. Panel C, A26 cells were surface-biotinylated at 4 °C and chased at 37 °C for the indicated times in the presence of the indicated concentrations of PS. Biotinylated chPrP was immunoprecipitated, run on SDS-PAGE, and blots of the gel developed using horseradish peroxidase-streptavidin and ECL. The amount of chPrP at each time point was quantitated from digitized film images and was expressed as percentage of the amount present immediately after biotinylation (0 h of chase). Each point represents the average of duplicate samples.



To determine whether PS affected the degradation rate of PrP, we pulse-labeled PrP and chased for up to 8 h in the presence or absence of 1 mg/ml PS. The half-life of PrP in both control and PS treated cells was 6 h (Fig. 5B). To selectively monitor the degradation rate of PrP residing on the cell surface, we quantitated the decrement in the amount of surface-biotinylated PrP during incubation in the presence or absence of PS. Again, no significant difference was found between PS-treated and control cells (Fig. 5C).

PS Increases the Amount of Intracellular PrP by Enhancing Endocytosis

Since it affected neither the synthesis nor degradation of PrP, PS was likely to reduce the amount of PrP on the cell surface by redistributing the protein to the cell interior. We initially investigated this possibility using a metabolic pulse-chase protocol. N2a cells were labeled for 30 min, and then chased for 30 min to allow newly synthesized molecules to reach the cell surface. After an additional chase for 1 h in the presence or absence of PS, cells were treated with PIPLC to separate surface from internal PrP. We found that PS shifted the distribution of PrP from the cell surface to an intracellular location (Fig. 6).


Figure 6: PS redistributes PrP from the surface to the interior of the cell. Panel A, A26 cells were labeled with TranS-Label for 30 min and chased for 30 min in the absence of PS. They were then chased for additional 60 min in the absence (Control) or presence (PS) of 1 mg/ml PS. Prior to lysis, cells were treated with PIPLC for 2 h at 4 °C to separate surface from internal chPrP. chPrP was immunoprecipitated from cell lysates (I lanes) and PIPLC incubation media (S lanes), and analyzed by SDS-PAGE and fluorography. Panel B, chPrP bands were quantitated using a PhosphorImager, and the amount of surface (S) and internal (I) chPrP was expressed as a percentage of the total amount of chPrP (surface+internal). Each bar shows the mean ± S.D. of values from 4 cultures. In the presence of PS, the total amount of chPrP was 110% of the amount in the absence of the drug.



We have previously shown that PrP^C constitutively cycles between the cell surface and an endocytic compartment in N2a cells (Shyng et al., 1993). To determine whether the observed shift in the localization of PrP resulted from an effect of PS on the rate of endocytosis, we directly measured internalization of surface-biotinylated PrP. We found that PS markedly enhanced the rate of PrP endocytosis after as little as 3 min (Fig. 7). PS does not affect the digestion efficiency of PIPLC, since >85% of surface-biotinylated chPrP was susceptible to digestion by the enzyme when cultures were maintained at 4 °C in either the presence or absence of the drug (data not shown). A similar stimulation of PrP endocytosis was also observed using lactoperoxidase-catalyzed iodination rather than biotinylation to label cells, and using trypsin rather than PIPLC to score the number of molecules remaining on the cell surface (data not shown).


Figure 7: PS stimulates endocytosis of PrP. Panel A, A26 cells were surface-biotinylated at 4 °C, and incubated at 37 °C for 3 min or 7 min in the absence (Cont) or presence (PS) of 1 mg/ml PS. Cells were then incubated with PIPLC for 2 h at 4 °C to separate cell surface from internal chPrP. chPrP was immunoprecipitated from cell lysates (C lanes) and PIPLC incubation media (M lanes), run on SDS-PAGE, and blots of the gel developed with horseradish peroxidase-streptavidin and ECL. Panel B, chPrP bands were quantitated from digitized film images, and the amount of internal chPrP was expressed as a percentage of the total amount of biotinylated chPrP (surface+internal) at each time point. Each bar shows the mean and range of values for duplicate cultures.



To determine whether the effect of PS on endocytosis is selective for PrP, we examined the rate of internalization of two other plasma membrane proteins. We first measured uptake of iodinated transferrin bound to its receptor, and found that PS had no significant effect on the percentage of prebound ligand that was internalized in 30 min (69.5 ± 4.8% and 66.7 ± 4.3% for PS-treated and control cells, respectively). We have previously reported that an N-terminally truncated form of chPrP, Delta25-91, is poorly endocytosed compared to wild-type chPrP (Shyng et al., 1995). In contrast to its dramatic effect on internalization of the wild-type protein, PS does not alter the normally low endocytic rate of Delta25-91 chPrP (12.5 ± 2.8% internalized after 30 min in PS-treated cells, compared to 12.6 ± 4.3% in control cells). In addition, overnight treatment with 1 mg/ml PS does not alter the steady-state level of Delta25-91 chPrP on the cell surface, as assayed by iodination with lactoperoxidase (not shown). These results suggest that the effect of PS is unlikely to be caused by a nonselective increase in the overall endocytic activity of the cell. The lack of effect on the Delta25-91 mutant further suggests that the N-terminal half of the chPrP molecule is important in mediating the activity of PS.

PS Redistributes Some PrP Molecules into Late Endosomes and/or Lysosomes

We performed electron microscopic immunogold labeling on ultrathin frozen sections to identify the intracellular compartments in which PrP accumulates after PS treatment. Cells were treated with anti-chPrP antibodies at 4 °C, and then incubated at 37 °C for 1 h in the presence or absence of PS to follow uptake of antibody-labeled molecules. After sectioning, samples were also labeled with antibodies to lgp120, a membrane protein that is a marker for late endosomes and lysosomes. Secondary antibodies coupled to gold particles of two different sizes were used to simultaneously visualize both antigens. In control cells, most of the chPrP molecules were found in vesicular structures that lacked lgp120, and are thus presumably early endosomes (Fig. 8A); this result is consistent with our previous immunofluorescence data (Shyng et al., 1993). In cells treated with PS, however, chPrP was also localized in some structures that were heavily labeled with anti-lgp120 antibodies (Fig. 8B).


Figure 8: PS redistributes some chPrP molecules into lgp120-positive compartments. A26 cells were incubated with anti-chPrP antibodies for 2 h at 4 °C and then chased for 1 h at 37 °C in the presence or absence of PS (1 mg/ml). Cells were then fixed and cryosectioned. Cryosections were incubated first with anti-lgp120 antibodies and then with secondary antibodies conjugated to 6- or 12-nm gold particles. In panels A and B, arrowheads point to 6-nm gold particles (chPrP), and arrows to 12-nm gold particles (lgp120). Panel A, a section from an untreated cell, showing chPrP and lgp120 labeling in distinct vesicles. Panel B, a section from a PS-treated cell, showing chPrP localization in vesicles that are also labeled for lgp120. Panel C, vesicles were categorized by the number of lgp120-associated gold particles they contained (horizontal axis), and the percentage of total chPrP-associated gold particles in each category of vesicles was calculated (vertical axis). Details of the quantitation are described under ``Materials and Methods.'' PS causes a statistically significant shift in the distribution of chPrP, such that more of the protein is localized in lgp120-positive vesicles, which represent late endosomes and/or lysosomes.



To quantitate the extent of co-localization of the two proteins, we categorized vesicles by the number of lgp120-associated gold particles they contained, and for each category counted the number of chPrP-associated gold particles (Fig. 8C). In control cells, 75% of the total PrP was found in compartments that were devoid of lgp120 staining, and only 7% in compartments that contained geq5 lgp120-associated particles. In PS-treated cells, in contrast, only 65% of the chPrP was in lgp120-negative structures, and 20% was found in structures having geq5 lgp-120 particles. The difference between the chPrP staining patterns in the presence and absence of PS was statistically significant (p < 0.005; ^2 test). These results indicate that PS causes a redistribution of some chPrP molecules to later compartments along the endocytic pathway, although the majority of the protein remains in early endosomal structures that do not contain lgp120.

A PrP-containing Bacterial Fusion Protein Binds to GAG Sites on Cultured Cells

Caughey et al.(1994) have shown that PrP^C binds in vitro to heparin and Congo red immobilized on beads, and they have postulated that the protein may also associate with GAG molecules synthesized by cells. To directly investigate the possible presence of PrP binding sites on cells, we used bacterially expressed fusion proteins that contain portions of the chPrP coding region fused to the E. coli maltose-binding protein (MBP). Binding to monolayers of N2a cells was assayed at 4 °C using iodinated MBP-chPrP fusion proteins. A fusion protein encompassing the mature chPrP polypeptide chain (residues 25-241) binds saturably to N2a cells, and binding is greatly reduced in the presence of excess unlabeled fusion protein (Fig. 9A). Scatchard analysis gives a K(d) of 230 nM and a capacity of 8.8 times 10^5 sites/cell (Fig. 9A, inset). A fusion protein encompassing the N-terminal half of the chPrP molecule (residues 25-116) binds to N2a cells with a similar affinity (240 nM), while a fusion protein encompassing the C-terminal half (residues 117-241) fails to bind (Fig. 9B). These results argue for the specificity of the binding reaction, and indicate that the N-terminal half of the polypeptide chain plays an essential role. There is a difference in the number of binding sites measured with the 25-241 and 25-116 fusion proteins (8.8 times 10^5/cell and 2.8 times 10^5/cell, respectively), suggesting that a subset of sites may require both the N- and C-terminal halves of the chPrP molecule for binding. We have also found that soluble chPrP released from brain membranes using PIPLC binds saturably and specifically to untransfected N2a cells, as determined using a chPrP-specific antibody (data not shown); this result indicates that native as well as bacterially produced protein binds to cultured cells.

We performed several experiments (data not shown) to further characterize the binding parameters for the 25-241 fusion protein. We found that binding was unaffected by the presence of 2 mM EDTA, arguing that it was not dependent on calcium. Binding was also not altered by variations in pH from 4 to 8. Finally, mild trypsinization of cells (0.05% for 10 min at 4 °C) reduced binding by almost 70%, arguing that a protein molecule is at least one component of the binding site.

We next carried out a series of experiments to investigate whether any of the binding sites for the 25-241 fusion protein might contain GAG chains. First, we found that rinsing N2a cells in 2 M NaCl, a treatment known to disrupt protein binding to GAGs (Moscatelli, 1987), removed over 80% of the fusion protein that had previously bound (Fig. 10A). Pretreatment of N2a cells with heparitinase, which digests heparan sulfate chains, reduced the amount of binding by 90% (Fig. 10A). In addition, exogenous GAGs effectively competed with the fusion protein for binding to N2a cells, with a rank order of potency PS > heparin > heparan sulfate > chondroitin sulfate (Fig. 10B). Finally, we analyzed binding of the 25-241 fusion protein to mutant CHO cells that lack GAGs because they are deficient in xylose transferase, the first enzyme required for GAG synthesis (Esko, 1991); these cells incorporate only 10% of the SO(4) label of wild-type cells (data not shown). The number of binding sites on the mutant CHO cells was 40% of the number on wild-type CHO cells, consistent with the idea that some, but not all of the sites are GAG molecules (Fig. 11). We noted that the K(d) for binding to wild-type and mutant CHO cells was similar and was lower than the K(d) for binding to N2a cells.


Figure 11: A xylose transferase-deficient CHO cell line has fewer binding sites than wild-type CHO cells for MBP-chPrP 25-241. Binding of iodinated fusion protein to wild-type (K1) and xylose transferase-deficient (clone 745) CHO cells was assayed as described in Fig. 9. Nonspecific binding, measured in the presence of 4 µM unlabeled fusion protein, was subtracted. Inset, Scatchard plots of the binding data shown in the main panel. Wild-type CHO cells (circles) show a K of 83 nM and 6.7 times 10^6 sites/cell; xylose transferase-deficient CHO cells (squares) show a K of 72 nM and 2.8 times 10^6 sites/cell.




DISCUSSION

Effect of Sulfated Anions on the Cellular Distribution of PrP^C

To understand the therapeutic effects of sulfated glycans and Congo red on prion production, we have examined the influence of these agents on the metabolism and cellular trafficking of PrP^C, the precursor of PrP. We report that PS, as well as Congo red and dextran sulfate, dramatically reduce the amount of PrP^C present on the cell surface. This effect results primarily from the ability of PS, and presumably the other agents, to stimulate endocytosis of PrP, thereby causing a redistribution of the protein from the plasma membrane to the cell interior. The effect is extremely rapid, occurring within 3 min of drug application. PS also causes a change in the ultrastructural localization of PrP, such that a portion of the protein molecules are shifted from early endosomes into late endosomes and/or lysosomes. Under the conditions tested, PS has no effect on the synthesis or degradation of PrP and does not change the net amount of the protein in the cell. Our results do not exclude the possibility that sulfated glycans produce other effects on cells when applied for longer periods than those used here. In this regard, Gabizon et al.(1993) reported that growth of N2a cells for 6 days in the presence of heparan sulfate increased the total cellular content of PrP.

The mechanism by which sulfated glycans induce such rapid and extensive alterations in the cellular distribution of PrP remains to be determined. These compounds appear to be relatively selective for PrP, since they affect both the chicken and mouse proteins but do not alter internalization of the transferrin receptor, or of a truncated form of chPrP. It therefore seems unlikely that the agents produce a general stimulation of endocytosis or of bulk membrane flow. However, an effect on other membrane trafficking events cannot be ruled out, since polyanions have been reported to influence processes such as phagosome-lysosomal fusion in macrophages (Hart and Young, 1975). We have demonstrated previously that PrP^C constitutively cycles between the cell surface and an endocytic compartment, and that internalization of PrP^C is mediated by clathrin-coated pits and vesicles (Shyng et al., 1993, 1994). The results reported here indicate that PS stimulates the endocytic arm of this cycle, although we do not yet know whether it is the clathrin pathway that is involved. The return arm of the cycle may also be affected, since PS inhibits iodinated PrP molecules from reappearing on the cell surface after they have been internalized (data not shown).

Sulfated glycans might act by binding directly to PrP on the cell surface, or by interacting with other cellular components that influence PrP trafficking. Consistent with a direct action, the N-terminal domains of chicken and mammalian PrP are rich in basic amino acids, and each contains a consensus site for heparin binding (XBBXBX, where B = basic amino acid, and X = other amino acids; Cardin and Weintraub, 1989); presumably, the same sites could mediate binding to other sulfated glycans. Moreover, we have found that a truncated form of chPrP (Delta25-91) that lacks this region is resistant to the endocytosis-promoting effect of PS. Binding of sulfated glycans to PrP is also consistent with our observation that chPrP fusion proteins bind to GAG-containing sites on the surface of cultured cells, and with the finding of Caughey et al.(1994) that PrP^C binds to heparin and Congo red immobilized on beads. How binding of sulfated glycans to cell surface PrP^C might enhance its endocytosis is a matter of speculation. One possibility is that these compounds induce oligomerization of PrP^C. It is well known that cross-linking of cell surface proteins by antibodies and other ligands rapidly stimulates their endocytosis (Marsh et al., 1995), and there are precedents for sulfated glycans causing oligomerization and aggregation of bound proteins (Jackson et al., 1991; Spivak-Kroizman et al., 1994).

It is possible that stimulation of PrP^C internalization by sulfated anions is related to the inhibitory effect of these agents on prion synthesis in cultured cells. Recent studies suggest that PrP^C is converted into PrP either on the cell surface, or along an endocytic pathway (Caughey et al., 1991b; Caughey and Raymond, 1991; Borchelt et al., 1992; Taraboulos et al., 1992). Sulfated glycans might therefore inhibit PrP production by promoting removal of PrP^C substrate from the plasma membrane, or by diverting PrP^C to an endocytic compartment that is unfavorable for the conversion process. Perhaps the lgp120-positive vesicles in which some of the PrP^C resides after PS treatment is such a compartment. We also note that the relative potencies of several different sulfated anions in reducing cell surface PrP are the same as their relative potencies in inhibiting PrP production (Fig. 4; Caughey and Raymond, 1993).

On the other hand, the dose of PS required for half-maximal removal of PrP^C from the cell surface (3-5 µg/ml; Fig. 3) is much higher than the dose required for half-maximal inhibition of PrP synthesis (1 ng/ml; Caughey and Raymond, 1993). This might imply that the two effects are unrelated. Indeed, Caughey et al.(1993) concluded that alterations in PrP^C metabolism did not underlie the scrapie-inhibitory effects of PS, based on their observation that treatment of N2a cells with 100 ng/ml PS did not affect the metabolic labeling, half-life, or PIPLC releasability of PrP^C during an 8.5 h chase. We have found, however, that even at 1 ng/ml, PS produced a 15-20% decrease of cell surface PrP^C after 24 or 48 h (Fig. 3), and it is possible that this effect is sufficient for inhibition of PrP production. It is also conceivable that PrP, rather than PrP^C, is the relevant target of PS action, and perhaps this isoform has a higher affinity for the drug. It will clearly be necessary to analyze the effects of sulfated glycans on the trafficking of PrP, as well as on other cellular processes relevant to prion synthesis, in order to fully understand the therapeutic action of these agents.

Cell Surface Binding Sites for PrP

The effect of exogenous glycans on membrane-bound PrP led us to search for structurally related GAG molecules on the cell surface that might bind to PrP. Using a radioiodinated fusion protein encompassing the mature chPrP sequence, we detected saturable and specific binding to monolayers of both N2a and CHO cells. Many of these binding sites are likely to be GAG molecules, as indicated by the measured dissociation constants, which are within the range seen for many GAG-protein interactions (Jackson et al., 1991), as well as by the inhibitory effects of 2 M NaCl, heparitinase, and exogenous GAGs, and by reduced binding to GAG-deficient CHO cells. In addition, a C-terminal PrP fragment that lacks consensus sites for heparin binding does not bind to cells. It is probable that the PrP binding sites on cultured cells comprise protein as well as GAG molecules, since treatment of cells with trypsin abolishes binding; a reasonable hypothesis is that the binding sites are proteoglycans. Caughey et al.(1994) have shown that a soluble form of PrP^C binds to Congo red and heparin when these compounds are immobilized on beads, but the work reported here is the first demonstration that GAG-containing binding sites for PrP exist on cells.

It is possible that membrane-associated molecules of PrP^C bind to cellular GAGs in a manner similar to that of the soluble PrP fusion proteins we have tested. If this were the case, then GAGs may play an important role in the function and metabolism of PrP^C. One potential function may involve endocytic trafficking of PrP^C. We have previously hypothesized the existence of a transmembrane receptor that binds glycosyl phosphatidylinositol-anchored molecules of PrP^C and mediates their internalization via clathrin-coated pits (Shyng et al., 1994, 1995; Harris et al., 1996). It is conceivable that binding to this putative receptor is facilitated by GAG chains, either those located on the receptor molecule itself or those added exogenously. Precedents for this model include the role of proteoglycans in endocytic uptake of several soluble ligands, and the role of heparin as an essential cofactor for binding of fibroblast growth factor to its high affinity receptor (Saxena et al., 1990; Klagsbrun and Baird, 1991; Reiland and Rapraeger, 1993; Rusnati et al., 1993; Kounnas et al., 1995). Other possible roles for the GAG binding sites we have identified include protecting surface PrP^C from extracellular proteases and trapping of soluble PrP^C that has been released by cleavage of the glycosyl phosphatidylinositol anchor.

GAG binding sites could potentially interact with PrP as well as PrP^C (Caughey and Raymond, 1993; Caughey et al., 1994; Priola and Caughey, 1994). To test this hypothesis, it will be interesting to directly assay binding of PrP to cell-associated GAGs and to assess the effect on prion synthesis of treatments that reduce GAG content, such as incubation with heparitinase or chlorate (Gabizon et al., 1993).

Although most of the binding sites for the PrP fusion proteins that we have detected appear to be GAGs, there is a suggestion that other chemically distinct sites may also exist. Even after extensive salt wash or heparitinase treatment, about 10-20% of the control binding always remains. In addition, xylose transferase-deficient CHO cells, which incorporate only 10% of the normal level of SO(4), display about half the number of binding sites as wild-type cells. These results suggest that PrP may also bind to non-GAG sites on cultured cells. The nature of these sites remains to be investigated.


FOOTNOTES

*
This work was supported by Grants NS30137 and AG12925 (to D. A. H.) from the National Institutes of Health. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
Recipient of fellowship support from National Institutes of Health Postdoctoral Training Grant T32 NS07071.

Recipient of a Postdoctoral Fellowship for Physicians from the Howard Hughes Medical Institute.

**
To whom all correspondence should be addressed: Dept. of Cell Biology and Physiology, Washington University School of Medicine, 660 S. Euclid Ave., St. Louis, MO 63110. Tel.: 314-362-4690; Fax: 314-362-7463; dharris@cellbio.wustl.edu.

(^1)
The abbreviations used are: PrP, scrapie isoform of the prion protein; CHO, Chinese hamster ovary; chPrP, chicken prion protein; ECL, enhanced chemiluminescence; GAG, glycosaminoglycan; PBS, phosphate-buffered saline; PIPLC, phosphatidylinositol-specific phospholipase C; PrP, prion protein; PrP^C, cellular isoform of the prion protein; PS, pentosan sulfate.


ACKNOWLEDGEMENTS

We thank Marilyn Levy and Lori LaRose for technical assistance with immunogold labeling of cryostat sections, and Jeffrey Esko for the gift of GAG-deficient CHO cells.


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