(Received for publication, July 5, 1995; and in revised form, August 7, 1995)
From the
Inherited forms of prion disease have been linked to mutations in the gene encoding PrP, a neuronal and glial protein that is attached to the plasma membrane by a glycosyl-phosphatidylinositol (GPI) anchor. One familial form of Creutzfeldt-Jakob disease is associated with a mutant PrP containing six additional octapeptide repeats. We report here our analysis of cultured Chinese hamster ovary cells expressing a murine homologue of this mutant PrP. We find that, like wild-type PrP, the mutant protein is glycosylated, GPI-anchored, and expressed on the cell surface. Surprisingly, however, cleavage of the GPI anchor using phosphatidylinositol-specific phospholipase C fails to release the mutant PrP from the surface of intact cells, suggesting that it has an additional mode of membrane attachment. The phospholipase-treated protein is hydrophobic, since it partitions into the detergent phase of Triton X-114 lysates; and it is tightly membrane-associated, since it is not extractable in carbonate buffer at pH 11.5. Whether membrane attachment of the mutant PrP involves integration of the polypeptide into the lipid bilayer, self-association, or binding to other membrane proteins remains to be determined. Our results suggest that alterations in the membrane association of PrP may be an important feature of prion diseases.
The spongiform encephalopathies are a group of transmissible
neurodegenerative disorders which are characterized by dementia, motor
dysfunction, and in some cases by the presence of cerebral amyloid
plaques (reviewed by Gajdusek, 1990; Prusiner and DeArmond, 1994).
These fatal disorders include Creutzfeld-Jakob disease (CJD), ()Gerstmann-Sträussler syndrome, kuru,
and fatal familial insomnia in human beings, and scrapie and bovine
spongiform encephalopathy in animals. The infectious agent responsible
for these diseases has been called a prion, to distinguish it from a
conventional virus (Prusiner, 1982). The major, and possibly only,
component of prion particles is a 35-45-kDa glycoprotein called
PrP
(Prusiner et al., 1984; Bolton et
al., 1987). PrP
is a post-translationally modified
isoform of a glycosyl-phosphatidylinositol (GPI)-anchored, cell surface
protein of the host called PrP
(Oesch et al.,
1985; Chesebro et al., 1985). Prion replication is
hypothesized to result from conversion of endogenous PrP
into infectious PrP
, a process that seems to involve
species-specific molecular interactions between the two isoforms
(Prusiner et al., 1990; Büeler et
al., 1993; Kocisko et al., 1994). Recent evidence
suggests that the isoforms differ in the conformation of their
polypeptide chains, and it has been proposed that conversion of
-helices into
-sheets underlies the formation of PrP
(Caughey et al., 1991; Safar et al., 1993; Pan et al., 1993). The two forms of the protein also exhibit
distinct biochemical properties. PrP
aggregates in
non-denaturing detergents, and it is more resistant to protease
digestion than PrP
, yielding an N terminally cleaved core
fragment called PrP 27-30 (Oesch et al., 1985; Meyer et al., 1986).
Prion diseases are unique in having both a genetic as well as an infectious origin. Gerstmann-Sträussler syndrome and fatal familial insomnia, and about 10% of the cases of CJD, are inherited in an autosomal dominant fashion (Prusiner and DeArmond, 1994). These inherited forms have been associated with specific mutations in the chromosomal gene that encodes PrP. Genetic linkage or association has been established for several amino acid substitutions in the C-terminal half of the molecule, as well as for insertions of between two and nine additional octapeptide repeats in the N-terminal half (reviewed by Brown et al., 1991; Prusiner and Hsiao, 1994). A genetic form of prion disease can also be produced in mice harboring a PrP transgene with a point mutation (Hsiao et al., 1990); mouse models of insertion mutations have not been reported.
It is presumed that
mutant PrPs spontaneously assume the conformation of PrP in the absence of exogenous prions (Prusiner and DeArmond, 1994).
However, there is no experimental evidence for this point, and the
cellular mechanisms that might be involved remain unknown. To address
these issues, it would be desirable to analyze the processing and
metabolism of mutant PrPs in cultured cells. We have previously
characterized the trafficking of wild-type PrP
in cultured
neuroblastoma cells and have defined the secretory and endocytic
pathways followed by the cellular isoform (Harris et al.,
1993; Shyng et al., 1993, 1994, and 1995). We have now
extended our studies to a murine form of PrP that carries an insertion
of six additional octapeptide repeats. This mutation is homologous to
one in human PrP that is strongly linked to CJD in a large family from
southeast England. This family comprises seven generations, and its
members are all descended from a single founder who was born in the
late 18th century (Owen et al., 1990; Poulter et al.,
1992; Collinge et al., 1992). Recently, two other families
with PrP insertions encoding the same amino acids have been published
(Nicholl et al., 1995; Oda et al., 1995). We report
here that murine PrP containing six additional octapeptide repeats
displays an abnormal association with the plasma membrane when
expressed in cultured CHO cells.
Figure 1: Structures of wild-type and PG11 moPrP constructs. Numbers beneath the schematics indicate amino acid positions. The numbers +1 through +6, above the PG11 schematic, indicate additional octapeptide repeats. The lollipop symbols indicate N-glycosylation sites, and GPI indicates the site of attachment of the glycolipid anchor. One set of constructs (untagged) had a leucine residue at wild-type position 108 (156 in PG11) and a valine residue at wild-type position 111 (159 in PG11). A second set of constructs (3F4-tagged) had methionine residues substituted at these positions, which allowed the proteins to be recognized by the monoclonal antibody 3F4.
To make PG11 moPrP, the following primers were first used to amplify the octapeptide repeat region of wild-type moPrP: 5`-GACCAGAAGCTTATGGCGAACCTTGGCTACTGG-3` (primer 3) and 5`-ATTATGGGTACCCCCTC-3` (primer 4). The amplified product was cleaved with BstXI, and a fragment of 72 base pairs comprising portions of the three central octapeptide repeats (R2-R3-R3 in Fig. 1) was isolated. This fragment was then ligated to itself, and after size fractionation of the ligation products on an agarose gel, a 216-base pair fragment was recovered. This fragment was then ligated to pBC12/CMV containing wild-type moPrP, after a 72-base pair fragment had been excised by digestion with BstXI.
To create moPrPs tagged with the 3F4 antibody epitope, the following PCR primers were employed: 5`-TGCCGCAGCCCCTGCCATATGCTTCATGTTGGTTTTTGGTTTGC-3` (primer 5), and 5`-GCAAACCAAAAACCAACATGAAGCATATGGCAGGGGCTGCGGCA-3` (primer 6). Untagged wild-type and PG11 cDNAs were first amplified with primer 1 and primer 5, and with primer 2 and primer 6. These two primary PCR products were then annealed with each other and amplified again with primer 1 and primer 2. After cleavage with HindIII and BamHI, the secondary PCR product was cloned into pBC12/CMV that had been cleaved with the same two enzymes.
The structure of each DNA construct was confirmed by sequencing. We note that moPrP contains a methionine residue at position 128, which is homologous to position 129 in human PrP where a valine/methionine polymorphism has been described; the PG11 gene in humans is always found to contain a methionine residue (Poulter et al., 1992).
N2a neuroblastoma cells expressing a chPrP/mannose 6-phosphate receptor (M6PR) transmembrane chimera are described elsewhere (Gorodinsky and Harris, 1995).
Samples from metabolically labeled cells were immunoprecipitated as described previously (Harris et al., 1993), with the modification that immunocomplexes were collected using protein A-Sepharose. Immunoprecipitated proteins were analyzed by SDS-PAGE, and radioactive gels were quantitated using a PhosphorImager (Molecular Dynamics).
Surface biotinylation was carried out as described previously (Shyng et al., 1995). Rabbit anti-mouse IgG was included when biotinylated proteins were immunoprecipitated with 3F4 antibody.
Figure 4:
Pulse-labeled molecules of both wild-type
and PG11 moPrP become resistant to endoglycosidase H and sensitive to
neuraminidase during the chase period. Panel A, CHO cells
expressing untagged wild-type (WT) or PG11 moPrP were labeled
with [S]methionine for 20 min and then chased in
medium containing unlabeled methionine for either 0 or 40 min. MoPrP
was immunoprecipitated from cell lysates using antibody P45-66
and was then either left untreated (- lanes) or was
digested with endoglycosidase H (+ lanes) prior to
SDS-PAGE. After 0 min of chase, both proteins show an increase in
electrophoretic mobility following endoglycosidase H treatment; this
shift is absent after 40 min of chase, indicating loss of high-mannose
type glycan chains. Panel B, CHO cells were labeled as in
panel A. MoPrP was immunoprecipitated from cell lysates using antibody
P45-66, and was then either left untreated (- lanes) or was digested with neuraminidase (+ lanes)
prior to SDS-PAGE. After 40 min of chase, both wild-type and PG11
moPrPs show an increase in electrophoretic mobility following treatment
with neuraminidase, indicating acquisition of sialic acid
residues.
In some experiments, we used epitopically tagged versions of wild-type and PG11 moPrP, in which methionine residues were substituted for a leucine and a valine residue at two nearby positions (Fig. 1). Methionine residues are found at the homologous positions in hamster and human PrP, and their introduction into moPrP allows the protein to be recognized by the species-specific monoclonal antibody 3F4 (Bolton et al., 1991). MoPrPs tagged with the 3F4 epitope have been used extensively in cell biological and transgenetic studies (Rogers et al., 1993; Scott et al., 1993).
We have expressed each of these moPrP molecules in stably transfected lines of CHO cells, which synthesize undetectable levels of endogenous hamster PrP (data not shown). A single cell line was analyzed for each construct. The unusual properties of PG11 moPrP that we describe here are not peculiar to a particular clone of cells, however, since both the untagged and 3F4-tagged versions of the mutant protein, expressed in separate lines, behaved similarly (see Fig. 6C). We noted that even though they had been subcloned, the lines expressing PG11 moPrP showed some decrease in their levels of PrP expression during prolonged passage. In most experiments, cells were used when the amounts of wild-type and mutant proteins were roughly equivalent (Fig. 2, Fig. 3, Fig. 4, Fig. 6, and Fig. 7). In two experiments ( Fig. 5and Fig. 8A), the amount of PG11 PrP was approximately 3-fold less than the amount of wild-type PrP. In none of our experiments, however, have we found that differences in expression levels had any effect on the results obtained.
Figure 6: Surface-biotinylated molecules of wild-type, but not PG11, moPrP are released by external PIPLC. Panel A, CHO cells expressing untagged wild-type (WT) or PG11 moPrP were biotinylated at 4 °C with the membrane-impermeant reagent sulfo-biotin-X-NHS. One set of cultures was then treated with PIPLC for 2 h at 4 °C prior to lysis, and moPrP in cell lysates (lanes 2 and 6) and PIPLC incubation medium (lanes 1 and 5) was analyzed by immunoprecipitation with antibody P45-66. Another set of cultures was treated with trypsin for 10 min at 4 °C (lanes 4 and 8), and a third set was left undigested (lanes 3 and 7), after which cells were lysed and moPrP immunoprecipitated. SDS-polyacrylamide gels of the immunoprecipitates were electroblotted, and blots were developed with horseradish peroxidase-streptavidin and ECL to visualize biotinylated PrP. Panel B, CHO cells expressing untagged wild-type (WT) or PG11 moPrP were biotinylated at 4 °C with sulfo-biotin-X-NHS and then treated with PIPLC at 4 °C for the indicated times. MoPrP in cell lysates and PIPLC incubation medium was then analyzed as described in panel A. PrP bands on the ECL film were quantitated by densitometry, and the amount of PrP released by PIPLC was plotted as a percentage of the total amount of PrP (medium + cell lysates). Each point represents the mean ± S.D. of three separate experiments. Panel C, CHO cells expressing 3F4-tagged wild-type (WT) or PG11 moPrP were biotinylated at 4 °C with sulfo-biotin-X-NHS and were then treated with PIPLC and processed as described in panel A, except that 3F4 was used as the primary antibody.
Figure 2: Wild-type and PG11 moPrPs are localized on the cell surface by immunofluorescence staining. CHO cells expressing untagged wild-type (A) or PG11 (B) moPrP, and untransfected CHO cells (C), were labeled with rabbit polyclonal antibody P45-66, and were then fixed with methanol, and stained with a fluorescein-coupled secondary antibody. Cells were observed by confocal microscopy. Each panel shows an optical section through the middle of the cells. Scale bar = 25 µm.
Figure 3:
Pulse-labeled molecules of both wild-type
and PG11 moPrP become accessible to external trypsin during the chase
period. Panel A, CHO cells expressing untagged wild-type (WT) or PG11 moPrP were labeled with
[S]methionine for 20 min and then chased in
medium containing unlabeled methionine for 0, 20, or 40 min. At the end
of the chase period, cells were either treated with trypsin for 10 min
at 4 °C (+ lanes) or remained untreated (- lanes) before lysis. MoPrP in cell lysates was then analyzed
by immunoprecipitation with antibody P45-66, either in the
presence (+ lanes) or absence (- lanes) of
the peptide immunogen. PrP-specific bands are those that are absent
when the peptide is included. Exposure time was 48 h for both gels. Panel B, PrP-specific bands in panel A were
quantitated by PhosphorImager analysis, and the amount of PrP digested
by trypsin at each time point was plotted as a percentage of the PrP
present in the absence of trypsin
treatment.
Figure 7:
The GPI anchors of both wild-type and PG11
moPrP incorporate [H]ethanolamine and
[
H]fatty acids and are cleavable by PIPLC. Panel A, CHO cells expressing untagged wild-type (WT)
or PG11 moPrP were labeled with either
[
S]methionine for 3 h, or with
[
H]ethanolamine for 16 h. Following enzymatic
deglycosylation, moPrP was immunoprecipitated from cell lysates using
antibody P45-66 and analyzed by SDS-PAGE. Arrowheads indicate PrP-specific bands. Panel B, CHO cells were
labeled with either [
S]methionine for 1 h (lanes 1-4) or with a mixture of
[
H]stearic acid and
[
H]palmitic acid for 16 h (lanes
5-8). Cells then either remained untreated (- lanes) or were incubated for 2 h at 4 °C with PIPLC (+ lanes) prior to lysis. MoPrP was then
immunoprecipitated from cell lysates after enzymatic deglycosylation
and analyzed by SDS-PAGE. Arrowheads indicate PrP-specific
bands. A small (1-2 kDa) decrease in the electrophoretic mobility
of PG11 PrP after cleavage with PIPLC can be seen by comparing lanes 3 and 4; this shift is more obvious in other
experiments using longer gel runs (data not
shown).
Figure 5:
Pulse-labeled molecules of wild-type, but
not PG11, moPrP are releasable by external PIPLC during the chase
period. Panel A, CHO cells expressing untagged wild-type (WT) or PG11 moPrP were labeled with
[S]methionine for 20 min and then chased in
medium containing unlabeled methionine for 0, 15, or 30 min. At the end
of the chase period, cells were treated with PIPLC for 2 h at 4 °C
before lysis. MoPrP in cell lysates (Cells) and PIPLC
incubation medium (Medium) was immunoprecipitated with
antibody P45-66 after treatment with N-glycosidase F to
remove N-linked oligosaccharides. A single band of PrP is
produced after deglycosylation (arrows). Exposure time was 48
h for both gels. Panel B, PrP bands in panel A were
quantitated by PhosphorImager analysis, and the amount of PrP released
by PIPLC was plotted as a percentage of the total amount of PrP present
at each time point (medium + cells). Panel C,
CHO cells expressing untagged wild-type (WT) or PG11 moPrP
were labeled with [
S]methionine for 4 h and were
then processed as described in panel A. PrP bands were
quantitated by PhosphorImager analysis, and the amount of PrP released
by PIPLC was plotted as a percentage of the total amount of PrP present (medium + cells). Each bar represents the mean
± S.D. of three separate
experiments.
Figure 8: PIPLC-treated PG11 PrP is retained in the detergent phase after Triton X-114 phase partitioning and is not extractable with alkaline carbonate buffer. Panel A, CHO cells expressing untagged wild-type (WT) or PG11 moPrP, and N2a cells expressing a chPrP/M6PR transmembrane chimera, were surface-biotinylated, and lysed at 4 °C in a buffer containing 1% Triton X-114. After the temperature was raised to 37 °C, the detergent phase was recovered, diluted to the original volume, and split in half. Half was incubated with PIPLC at 4 °C for 2 h (+ lanes), and the other half was left untreated (- lanes). Phase separation was then repeated, and moPrP in the detergent (D) and aqueous (A) phases was immunoprecipitated with antibody P45-66 (lanes 1-8); chPrP/M6PR was immunoprecipitated with antibody F35-96 (lanes 9-12). SDS-polyacrylamide gels of the immunoprecipitates were electroblotted, and the blots developed with horseradish peroxidase-streptavidin and ECL to visualize biotinylated proteins. Panel B, PrP bands from the experiment shown in panel A, and from two additional experiments, were quantitated by densitometry, and the amount of PrP in the detergent phase after PIPLC treatment (lanes 3, 7, and 11) was plotted as a percentage of the total amount of PrP present (detergent + aqueous phases). Each bar represents the mean ± S.D. Panel C, CHO cells expressing PG11 moPrP were surface-biotinylated and incubated with PIPLC for 2 h at 4 °C. Cells were then homogenized in PBS, and a postnuclear membrane fraction prepared. Membranes were resuspended either in PBS or in carbonate buffer and were then collected by centrifugation. MoPrP in the pellet and supernatant fractions was then analyzed by immunoprecipitation, followed by SDS-PAGE, electroblotting, and visualization with HRP-streptavidin and ECL (lanes 1-4; bracket indicates PrP-specific bands). Pellet and supernatant fractions were also immunoblotted to detect clathrin heavy chain (arrow), a marker peripheral membrane protein (lanes 5-8).
These results suggested that the PG11 protein reached the cell surface after synthesis, in contrast to some other mutant proteins which remain trapped intracellularly as a result of aberrant folding of the polypeptide chain in the endoplasmic reticulum (Amara et al., 1992). To directly examine the kinetics with which the mutant protein was delivered to the cell surface, we performed a metabolic pulse-chase experiment (Fig. 3). We quantitated the amount of PrP transported to the cell surface by determining the proportion of labeled protein that was digested after incubation of intact cells with trypsin. In this experiment, we found that both wild-type and PG11 forms of moPrP were transported to the surface with similar kinetics (Fig. 3B). The two proteins both became accessible to external trypsin by 20 min of chase, and by 40 min approximately 60% was susceptible to digestion by the enzyme. Similar kinetics have been reported for delivery of pulse-labeled molecules of chicken PrP (Shyng et al., 1993) and moPrP (Caughey et al., 1989; Borchelt et al., 1992) to the surface of mouse neuroblastoma cells.
To further explore the glycosylation of PG11 moPrP, we have tested the susceptibility of the protein to digestion by endoglycosidase H and neuraminidase after pulse-labeling. During the chase period, both wild-type and PG11 PrPs become resistant to digestion by endoglycosidase H (Fig. 4A) and sensitive to digestion by neuraminidase (Fig. 4B). These results indicate that both proteins traverse the mid-Golgi stack (site of acqui-sition of endoglycosidase H resistance) and the trans-Golgi network (site of sialic acid addition) during their passage to the surface of CHO cells. Maturation of the oligosaccharide chains of moPrP to an endoglycosidase H-resistant form has also been observed in neuroblastoma cells (Caughey et al., 1989; Taraboulos et al., 1992). Taken together, our results indicate that the presence of additional octapeptide repeats does not markedly affect the kinetics or route of biosynthetic processing of PrP, but does affect the specific pattern of glycosylation.
We first examined this question by performing a pulse-chase labeling experiment similar to that shown in Fig. 3, but using PIPLC rather than trypsin (Fig. 5). Cells were incubated with PIPLC at each time point and then lysed; the amount of PrP in cell lysates and PIPLC incubation media was determined by immunoprecipitation. During the chase period, wild-type moPrP gradually became accessible to digestion by PIPLC, so that by 30 min almost 40% of the protein was recovered in the PIPLC incubation medium (Fig. 5B). This increase reflects delivery of newly synthesized molecules to the cell surface, where they were susceptible to release by the phospholipase. The kinetics of surface delivery of wild-type moPrP measured in this way was quite similar to that measured using external trypsin (compare Fig. 3B and Fig. 5B).
Surprisingly,
a very different result was obtained with CHO cells expressing the PG11
form of moPrP. Even after 30 min of chase, very little of the mutant
protein was released by PIPLC, and most of it remained associated with
the cell lysates (Fig. 5B). A similar result was
obtained when cells were labeled continuously for 4 h with
[S]methionine, eliminating the possibility that
PG11 molecules become releasable by PIPLC at later times (Fig. 5C). The fact that PG11 moPrP cannot be released
by the PIPLC contrasts with our observation that almost half of the
mutant protein is susceptible to digestion by external trypsin 30 min
after pulse labeling (Fig. 3B). Taken together, these
data indicate that the PG11 protein is not releasable by PIPLC, even
though substantial amounts of it are present on the cell surface.
To
confirm this conclusion, we analyzed the effect of PIPLC on cell
surface PrP that had been labeled by biotinylation of intact cells with
the membrane-impermeant reagent sulfo-biotin-X-NHS. We found that
90% of wild-type moPrP was released by PIPLC, in contrast to
<5% of PG11 moPrP (Fig. 6A, lanes 1, 2, 5, and 6). As a control to confirm that
only PrP molecules on the cell surface had been biotinylated, we found
that both forms of PrP were completely digested by externally applied
trypsin (Fig. 6A, lanes 3, 4, 7, and 8). Lack of release of PG11 did not result
from a slower action of PIPLC on the mutant protein, since extending
incubation with the phospholipase to as long as 6 h failed to liberate
any additional PrP (Fig. 6B). We obtained similar
results after surface iodination of cells, indicating that the failure
of PIPLC to release PG11 is not an artifact of biotinylation (data not
shown).
To confirm that the unusual behavior of the mutant protein
was not a peculiarity of the particular clone of CHO cells we were
analyzing, we performed an identical biotinylation experiment on a
separate pair of cell lines that expressed moPrPs tagged with the 3F4
antibody epitope (Fig. 6C). In agreement with our
previous results, we found that >90% of wild-type moPrP was released
by PIPLC, in contrast to 5% of the PG11 mutant. We noted in this
and other experiments that the small amount of the mutant protein
released was usually the most heavily glycosylated isoform (Fig. 6C, lane 3). In preliminary experiments,
we have also observed that PIPLC fails to release human PG11 PrP from
the surface of biotinylated CHO cells (data not shown). We have also
found that similar results are obtained whether CHO cells are stably or
transiently transfected (not shown).
Our data thus far
indicated that PG11 moPrP molecules are GPI-anchored, even though they
cannot be released from the cell surface by digestion with PIPLC. A
possible explanation for this fact might be that the mutant protein
contains a modified GPI anchor that is intrinsically resistant to PIPLC
cleavage, as has been reported for anchor structures in which the
inositol ring is acylated (Rosenberry, 1991). Alternatively,
aggregation of the protein on the cell surface, or interaction with
other membrane components, might render the anchor physically
inaccessible to the phospholipase. To investigate these possibilities,
we incubated CHO cells with a mixture of
[H]palmitate and
[
H]stearate in order to label the fatty acyl
residues of the GPI anchor and then treated the intact cells with
PIPLC. Since PIPLC cleaves the diacylglycerol moiety from the GPI
anchor, the activity of this enzyme would be revealed by a loss of
incorporated
H label after PIPLC treatment. We also labeled
parallel cultures with [
S]methionine to track
the polypeptide chain. We found that both wild-type and PG11 moPrP
molecules were labeled with
H-fatty acids, consistent with
the presence of a GPI anchor on both proteins (Fig. 7B, lanes 5 and 7). Importantly, PIPLC treatment of the
cells removed the
H label from the PG11 protein almost
completely (Fig. 7B, lane 8). Analysis of the
[
S]methionine-labeled dishes confirmed that
after removal of the diacylglycerol moiety by PIPLC, PG11 molecules
were retained on the cell surface (Fig. 7B, lane
4), while the wild-type molecules were released into the medium (Fig. 7B, lane 2). This result demonstrates
that the GPI anchor of the PG11 mutant is cleavable by PIPLC, but that
cleaved molecules are not released from the cell surface. Consistent
with this conclusion, we also observed a small (1-2 kDa) decrease
in SDS-PAGE mobility of the PG11 protein after PIPLC treatment (Fig. 7B, lanes 3 and 4). This shift
in mobility is characteristic of many GPI-anchored proteins after
digestion with PIPLC and is probably due to changes in SDS binding
after removal of diacylglycerol (Ferguson and Williams, 1988;
Rosenberry, 1991).
First, we subjected moPrP to phase partitioning in the detergent Triton X-114. In this method, integral membrane and other hydrophobic proteins are found in the detergent phase, while peripheral membrane and soluble proteins are found in the aqueous phase (Bordier, 1981). We found that before treatment with PIPLC, both wild-type and PG11 moPrPs partitioned almost exclusively into the detergent phase, as would be expected for molecules carrying an intact GPI anchor (Fig. 8A, lanes 1 and 2 and 5 and 6). PIPLC treatment shifted the wild-type protein almost completely into the aqueous phase (Fig. 8A, lanes 3 and 4 and Fig. 8B), a phenomenon that is observed for other GPI-anchored proteins (Englund, 1993), and which results from loss of the hydrophobic diacylglycerol portion of the anchor. In contrast, the mutant protein was only partially shifted, with over half remaining in the detergent phase after PIPLC treatment (Fig. 8A, lanes 7 and 8 and Fig. 8B). As a control, we analyzed the phase-partitioning of a transmembrane protein, a chPrP/mannose 6-phosphate receptor chimera (Gorodinsky et al., 1995), and found that it was almost completely retained in the detergent phase, whether or not it was treated with PIPLC (Fig. 8A, lanes 9-12 and Fig. 8B). We thus conclude that, even after removal of the diacylglycerol portion of the GPI anchor, the PG11 molecule displays substantial hydrophobicity.
As an additional test of the nature of the association between PG11 and the plasma membrane, we determined whether the protein was extractable by carbonate buffer at pH 11.5. Because many protein-protein interactions are disrupted at this pH, carbonate inextractability is commonly used as a criterion for stable integration of a polypeptide into the lipid bilayer (Fujiki et al., 1982). We prepared a membrane fraction from biotinylated CHO cells that had been treated with PIPLC and incubated it on ice with either PBS or carbonate buffer. We then collected the membranes by centrifugation and assayed the amount of PrP in the pellet and supernatant. We found that, even after PIPLC treatment, very little of the PG11 protein was extractable from the membranes by carbonate buffer (Fig. 8C, lanes 3 and 4), consistent with an alkaline-stable association. As a control for the effectiveness of the extraction conditions, we immunoblotted the supernatant and pellet fractions using an antibody to the heavy chain of clathrin and found that the majority of this peripheral membrane protein was solubilized by carbonate buffer, but not by PBS (Fig. 8C, lanes 5-8).
The data reported here demonstrate that a mutant form of moPrP containing 11 octapeptide repeats displays an aberrant mechanism of membrane association when expressed in cultured CHO cells. These results are directly relevant to human prion diseases, since an insertional mutation in the PrP gene that results in 11 octapeptide repeats has been strongly linked to familial CJD in a large English family (Poulter et al., 1992; Collinge et al., 1992) and has also been described in two smaller families (Nicholl et al., 1995; Oda et al., 1995). This report presents the first description of an abnormality in the biochemical properties of a mutant PrP molecule, and it raises the possibility that an alteration in the membrane attachment of PrP is a critical feature of prion diseases.
We have presented several lines of evidence indicating
that, while both wild-type and PG11 forms of PrP are glycosylated and
delivered to the cell surface, they differ dramatically in their mode
of membrane association. The surface localization of both proteins in
intact cells has been shown by immunofluorescence staining, as well as
by accessibility to externally applied biotinylation and iodination
reagents, and to trypsin. Both wild-type and PG11 PrPs also contain a
GPI anchor, as demonstrated by metabolic incorporation of
[H]ethanolamine and
H-fatty acids,
which are anchor constituents. The critical difference in membrane
attachment between wild-type and PG11 PrP is revealed by digestion of
intact cells with PIPLC, which is a bacterial enzyme that specifically
cleaves the GPI anchor, removing the diacylglycerol portion that is
embedded in the lipid bilayer. While the wild-type protein is released
nearly quantitatively by PIPLC, the PG11 protein is almost completely
retained on the cell surface after digestion. We have shown that the
inability of PIPLC to release PG11 PrP is not due to a failure to
cleave the GPI anchor, since the phospholipase almost completely
removes
H-fatty acid label from the protein when applied to
intact cells, and produces a characteristic decrease in its
electrophoretic mobility. These observations make it unlikely that
aggregation of the mutant protein on the cell surface, or interaction
with other membrane components, renders the GPI anchor inaccessible to
enzymatic cleavage, at least for the majority of the molecules;
however, our data do not rule out the possibility that the anchors of a
small proportion of the molecules are resistant to cleavage.
The results indicate that PG11 PrP must have a second mechanism of membrane attachment in addition to the GPI anchor. Several possible models can be envisioned. One is that the PG11 polypeptide chain is directly integrated into the lipid bilayer. In support of this proposal, we find that after cleavage of the GPI anchor with PIPLC, 65% of the mutant PrP still partitions into the detergent phase after Triton X-114 phase separation (Bordier, 1981), in comparison to 15% for wild-type PrP (Fig. 8B). These results indicate that the PG11 molecule is quite hydrophobic, even after removal of diacylglycerol from the GPI anchor, suggesting that a portion of the protein has the potential to interact with lipids. The fact that 85% of a bona fide transmembrane protein was retained in the detergent phase after partitioning in Triton X-114 (Fig. 8B) might indicate that PG11 possesses a shorter or less hydrophobic lipid-associated domain than the transmembrane protein. Direct association between the polypeptide chain and the lipid bilayer is also suggested by failure to release PIPLC-digested PG11 from isolated membranes using alkaline carbonate buffer, which disrupts many protein-protein interactions. We note that transmembrane forms of PrP have been observed when PrP mRNA is translated in cell-free systems (Hay et al., 1987; Yost et al., 1990; Lopez et al., 1990; De Fea et al., 1994) and that several other proteins are proposed to have both a GPI anchor and a membrane-integrated polypeptide chain (Köster and Strand, 1994; Hitt et al., 1994; Howell et al., 1994). Nevertheless, it will be necessary to employ additional techniques, such as labeling with lipid-soluble reagents and mapping the accessibility of sequence epitopes, to decide whether or not the PG11 polypeptide chain loops into or traverses the lipid bilayer.
A second possible model is that PG11 moPrP binds
tightly to other membrane-associated molecules, thereby preventing
release of mutant PrP after cleavage of its PIPLC anchor. This
hypothetical interaction would have to be stable in carbonate buffer at
pH 11.5. One potentially relevant class of surface binding sites is
proteoglycan molecules. PrP binds to heparin-agarose beads
(Caughey et al., 1994), and we have found that PrP-containing
bacterial fusion proteins bind to glycosaminoglycan sites on the
surface of cultured cells. (
)In this model, one would have
to postulate that the conformation of the PG11 molecule is altered in
such a way that it binds more tightly than the wild type molecule to
membrane-associated sites.
A final possibility is that PG11 moPrP
forms aggregates on the cell surface and that these remain attached to
the membrane after PIPLC treatment by virtue of a small number of
molecules whose GPI anchors have escaped, or are resistant to,
cleavage. In fact, we have found the PG11 protein expressed in CHO
cells forms detergent-insoluble aggregates that can be recovered by
ultracentrifugation. ()Arguing against an effect of
aggregation, however, is the high efficiency of PIPLC cleavage
(
90%, as judged by loss of
H-fatty acid label; Fig. 7B, lanes 7 and 8); and the
observation that no additional PG11 protein is released by prolonging
the period of PIPLC digestion to 6 h (Fig. 6B).
We
have noted that the glycosylation pattern of PG11 PrP differs from that
of wild-type PrP, with the mutant protein displaying a larger
proportion of less glycosylated forms (Fig. 6A). We
have also observed that the small amount (1-5%) of PG11 PrP that
can be released by PIPLC is usually the most heavily glycosylated form (Fig. 6C). These results suggest that there may be a
relationship between the altered glycosylation state of mutant PrP and
its aberrant membrane association, a proposal consistent with the
observation that PrP synthesized in infected neuroblastoma
cells also displays a predominance of less glycosylated forms (Caughey
and Raymond, 1991).
We have recently found that PG11 moPrP produced
in CHO cells displays two of the characteristic biochemical properties
of PrP, aggregation in non-ionic detergent, and resistance
to protease digestion.
We have not yet tested whether the
protein is infectious. It will be important now to determine how these
scrapie-like properties are related to the abnormal membrane
association of PG11, and whether the conformational changes that
underlie generation of PrP
involve alterations in membrane
attachment. Although there are no data available on the protease
sensitivity of PG11 PrP from human patients, these individuals do not
develop symptoms until adulthood (Collinge et al., 1992),
suggesting that accumulation of PrP
is a slow process that
may require many years. Since we find that PG11 moPrP molecules in CHO
cells acquire an aberrant membrane association, and become
protease-resistant and detergent-insoluble,
within a matter
of hours, the generation of PrP
may be more efficient in
CHO cells than in neurons in vivo. There is one report that
CHO cells cannot be infected by scrapie prions (Butler et al.,
1988), but the cellular factors that influence prion replication are
poorly understood and may be different from those that affect
PrP
production from mutant molecules.
Although our
study has focused on a mutant form of PrP, our results lead us to
hypothesize that the PrP produced during prion infection
possesses an aberrant membrane association similar to that of PG11.
This proposal is generally consistent with a large body of literature,
some of it dating back more than 30 years, suggesting that infectious
PrP
is tightly associated with cellular membranes, and is
very hydrophobic (Hunter et al., 1964; Gibbons and Hunter,
1967; Prusiner et al., 1978). Of particular relevance, it has
been reported that, although PrP
possesses a GPI anchor,
it is not readily releasable from brain membranes by PIPLC (Stahl et al., 1990; Safar et al., 1991). Lack of release by
PIPLC does not result from sequestration in a membrane-bound
compartment, since PrP
is susceptible to biotinylation
with membrane-impermeant reagents, and to cleavage by proteases;
moreover, the GPI anchor of purified PrP
is
PIPLC-sensitive, arguing that the anchor is not intrinsically
uncleavable. These properties of PrP
from infected brain
are strikingly reminiscent of those we have described for PG11 PrP in
CHO cells. It is uncertain from the published studies whether the
failure of PIPLC to release membrane-bound PrP
results
from inaccessibility of the anchor structure, or rather, signifies a
second, GPI-independent form of membrane attachment. Our results with
PG11 PrP argue for the second possibility. It will be useful now to
reexamine the membrane association of PrP
from
scrapie-infected cells in culture, using experimental approaches
similar to those we have described here.
Most of the known
pathogenic mutations in human PrP are single amino acid substitutions
in the C-terminal half of the molecule, in contrast to the 48 amino
acid insertion in the N-terminal half that we have modeled here
(Prusiner and Hsiao, 1994). We find that several moPrPs carrying point
mutations homologous to those found in human PrP are not released from
the surface of transfected CHO cells by PIPLC, implying an aberrant
form of membrane association like that described for PG11. ()These data support the idea that alterations in membrane
attachment may be a general feature of both genetic and infectious
forms of prion disease.