(Received for publication, August 7, 1995; and in revised form, October 5, 1995)
From the
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
, 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
,
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
, 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
precursor, and they indicate that endogenous
proteoglycans are likely to play an important role in the cellular
metabolism of both PrP
and PrP
.
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(
)(Prusiner et
al., 1984; Bolton et al., 1987). PrP
is a
posttranslationally modified isoform of PrP
, 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
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
-helices into
-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
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
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
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 or PrP
might alter the efficiency of prion
production. Recent studies of scrapie-infected neuroblastoma cells
suggest that conversion of PrP
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
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
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
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, 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
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
precursor,
and they suggest that endogenous proteoglycans are likely to play an
important role in the cellular metabolism of both PrP
and
PrP
.
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).
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.
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.
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
10
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
10
sites/cell.
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.
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 Tran
S-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).
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 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, 25-91, 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
25-91
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
25-91 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
25-91 mutant further suggests that the N-terminal half of
the chPrP molecule is important in mediating the activity of PS.
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 5 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
5 lgp-120 particles. The difference between the chPrP staining
patterns in the presence and absence of PS was statistically
significant (p < 0.005;
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.
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
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
for binding to wild-type and mutant CHO cells was similar and was
lower than the K
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
10
sites/cell; xylose transferase-deficient CHO cells (squares) show a K
of 72 nM and 2.8
10
sites/cell.
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 constitutively cycles between the cell surface and
an endocytic compartment, and that internalization of PrP
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 (25-91) 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
binds to
heparin and Congo red immobilized on beads. How binding of sulfated
glycans to cell surface PrP
might enhance its endocytosis
is a matter of speculation. One possibility is that these compounds
induce oligomerization of PrP
. 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 internalization by sulfated anions is
related to the inhibitory effect of these agents on prion synthesis in
cultured cells. Recent studies suggest that PrP
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
substrate from
the plasma membrane, or by diverting PrP
to an endocytic
compartment that is unfavorable for the conversion process. Perhaps the
lgp120-positive vesicles in which some of the PrP
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 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
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
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
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
, 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.
It is possible
that membrane-associated molecules of PrP 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
. One potential
function may involve endocytic trafficking of PrP
. We have
previously hypothesized the existence of a transmembrane receptor that
binds glycosyl phosphatidylinositol-anchored molecules of PrP
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
from extracellular proteases and trapping of
soluble PrP
that has been released by cleavage of the
glycosyl phosphatidylinositol anchor.
GAG binding sites could
potentially interact with PrP as well as PrP
(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
, 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.