(Received for publication, January 14, 1997, and in revised form, February 18, 1997)
From the Department of Cell Biology and Physiology, Washington University School of Medicine, St. Louis, Missouri 63110
The central causative event in infectious, familial, and sporadic forms of prion disease is thought to be a conformational change that converts the cellular isoform of the prion protein (PrPC) into the scrapie isoform (PrPSc) that is the primary constituent of infectious prion particles. To provide a model system for analyzing the mechanistic details of this critical transformation, we have previously prepared cultured Chinese hamster ovary cells that stably express mouse PrP molecules carrying mutations homologous to those seen in familial prion diseases of humans. In the present work, we have analyzed the kinetics with which a PrP molecule containing an insertional mutation associated with Creutzfeldt-Jakob disease acquires several biochemical properties characteristic of PrPSc. Within 10 min of pulse labeling, the mutant protein undergoes a molecular alteration that is detectable by a change in Triton X-114 phase partitioning and phenyl-Sepharose binding. After 30 min of labeling, a detergent-insoluble and protease-sensitive form of the protein appears. After a chase period of several hours, the protein becomes protease-resistant. Incubation of cells at 18 °C or treatment with brefeldin A inhibits acquisition of detergent insolubility and protease resistance but does not affect Triton X-114 partitioning and phenyl-Sepharose binding. Our results support a model in which conversion of mutant PrPs to a PrPSc-like state proceeds in a stepwise fashion via a series of identifiable biochemical intermediates, with the earliest step occurring during or very soon after synthesis of the polypeptide in the endoplasmic reticulum.
Prion diseases are a group of unusual neurodegenerative disorders that includes Creutzfeldt-Jakob disease (CJD),1 kuru, Gerstmann-Sträussler syndrome, and fatal familial insomnia in human beings, as well as scrapie and bovine spongiform encephalopathy in animals (reviewed in Ref. 1). These disorders are characterized clinically by dementia and motor dysfunction and neuropathologically by spongiform degeneration of the brain and in some cases by the presence of cerebral amyloid plaques. Prion diseases can have an infectious, familial, or sporadic origin. The infectious etiology is exemplified by kuru, which was spread among members of an aboriginal tribe by ritual cannibalism (2), and by "new variant" CJD which may have arisen by consumption of beef from cows afflicted with bovine spongiform encephalopathy (3). Familial forms, which include Gerstmann-Sträussler syndrome, fatal familial insomnia, and 10% of the cases of CJD, show an autosomal dominant mode of inheritance linked to insertional and point mutations in the gene encoding the prion protein (PrP) (4).
The central pathogenic event in prion diseases is thought to be a
conformational transition that changes PrPC, a normal cell
surface protein of unknown function, into PrPSc, the
principal component of infectious prion particles (1, 5). This change
is accompanied by transformation of -helices into
-sheets in
critical sections of the polypeptide chain (6-8). However, many
mechanistic details of the conversion process remain obscure. First,
the nature of the interaction between PrPSc and
PrPC is poorly understood. A physical association between
the two isoforms during the infectious process is suggested by the
exquisite primary sequence specificity of prion transmission (9-11)
and by the recent demonstration that PrPSc-like molecules
can be generated in vitro by mixing purified
PrPC with PrPSc or with synthetic PrP peptides
(12, 13). However, a PrPC-PrPSc conversion
intermediate has never been isolated from cells, and there is
considerable debate about the whether the conversion process is best
modeled as a nucleated polymerization (12, 14-16), a
chaperone-assisted refolding (17), or some combination of the two
mechanisms. Second, it is unclear how the hypothesized change in PrP
conformation alters the experimentally observed properties of the
protein. Whether the conversion of PrPC to infectious
PrPSc occurs in an all-or-nothing fashion or proceeds via a
series of identifiable molecular intermediates is unknown. Third, there is uncertainty about the cellular compartments involved in generation of PrPSc. Evidence from scrapie-infected cells in culture
implicates the plasma membrane, endocytic organelles, as well as
cholesterol-rich microdomains as relevant sites (18-21), but it is
unclear which of these is most important and whether compartments along
the secretory pathway might also be involved. Finally, it remains to be
determined what molecules other than PrP itself might be important in
the conversion process. Although structural features intrinsic to PrP
probably play a critical role in the prion phenomenon, cellular
cofactors may function to enhance or modulate PrPSc
formation, as suggested by the inefficiency of existing in
vitro conversion systems (12, 13), the well known role of
chaperone molecules in protein folding (22), and by recent transgenic experiments that implicate a hypothetical protein "X" in
prion replication (23).
To address these mechanistic issues, it is necessary to analyze prion formation in cultured cells that are amenable to experimental manipulations such as pulse labeling, microscopy, and subcellular fractionation. Scrapie-infected neuroblastoma cells and immortalized hamster brain cells have been invaluable in elucidating some of the cellular and biochemical events underlying the infectious manifestation of prion diseases (18-21, 24, 25). Until recently, however, there has been no cell culture model of familial forms of prion disease. We (26-28) and others (29) have now developed such a model. Our system utilizes transfected lines of Chinese hamster ovary (CHO) cells that stably express mouse PrP (moPrP) molecules carrying mutations homologous to those linked to inherited prion diseases of humans. We find that mutant PrPs synthesized in these cells display many of the biochemical properties of PrPSc, including detergent insolubility, protease resistance, slow metabolic generation and turnover, abnormally tight membrane attachment, and strain-like variations in glycosylation pattern and protease-cleavage site. We have now used this model system to examine the kinetics with which mutant PrP molecules acquire PrPSc-like properties and to draw inferences from this about the nature of the intermediates in the conversion process and the identity of the cellular compartments in which they are generated.
Cell culture reagents were from the Tissue Culture Support Center at Washington University. N-Glycosidase F was purchased from Boehringer Mannheim, sulfo-biotin-X-NHS was from Calbiochem, brefeldin A (BFA) was from Sigma, phenyl-Sepharose CL-4B was from Pharmacia Biotech Inc., and [35S]methionine (Pro-mix, 1,000 Ci/mmol) was from Amersham Corp. Phosphatidylinositol-specific phospholipase C (PIPLC) from Bacillus thuringiensis was prepared as described previously (30).
Rabbit polyclonal antibody P45-66, raised against a synthetic peptide encompassing moPrP residues 45-66, has been described (26). Rabbit polyclonal anti-ME7 and mouse monoclonal 3F4 antibodies, raised against PrP 27-30 from scrapie-infected mouse and hamster brain, respectively, were gifts from Dr. Rick Kascsak (31). Although the latter two antibodies react with hamster PrP, CHO cells do not synthesize detectable levels of endogenous hamster PrP,2 so that only recombinant moPrP is detected in this cell type.
Cell LinesStably transfected lines of CHO cells expressing wild-type, PG11, and E199K moPrPs have been described previously (26-28). The wild-type and PG11 constructs contained an epitope tag for the monoclonal antibody 3F4.
Assay of Detergent InsolubilityConfluent cultures of CHO cells were labeled in methionine-free minimal essential medium containing [35S]methionine (Pro-mix, 250-500 µCi/ml) and were chased in Opti-MEM (Life Technologies, Inc.). Cells were then lysed in a buffer that contained 150 mM NaCl, 50 mM Tris-HCl (pH 7.5), 0.5% Triton X-100, and 0.5% sodium deoxycholate, supplemented with protease inhibitors (pepstatin and leupeptin, 1 µg/ml; phenylmethylsulfonyl fluoride, 0.5 mM; EDTA, 2 mM). Lysates were first centrifuged for 5 min at 16,000 × g in a microcentrifuge, a procedure that removes debris but does not pellet significant amounts of PrP. The cleared lysates were then centrifuged at 265,000 × g for 40 min in the TLA 100.3 rotor of a Beckman Optima TL ultracentrifuge to separate detergent-soluble and detergent-insoluble protein. Immunoprecipitation of moPrP in pellet and supernatant fractions from the second centrifugation was performed using monoclonal antibody 3F4 antibody as described previously (26). In some experiments lysates were treated with N-glycosidase F (0.01 units/ml) for 16 h at 37 °C prior to immunoprecipitation to produce a single band of deglycosylated PrP that could be more easily quantitated (32). Immunoprecipitated proteins were analyzed by SDS-polyacrylamide gel electrophoresis (PAGE), and radioactive gels were quantitated using a PhosphorImager (Molecular Dynamics).
Assay of Protease ResistanceCells were metabolically labeled and chased as above. Proteins were methanol-precipitated from cell lysates, digested with proteinase K (3.3 µg/ml for 10 min at 37 °C) in Sarkosyl, methanol-precipitated a second time, and then dispersed in detergent-lipid-protein complexes for immunoprecipitation of moPrP using anti-ME7 antibody, all as described (19, 27, 28). Immunoprecipitated proteins were analyzed by SDS-PAGE.
For the experiment shown in Fig. 1C, lysates of unlabeled
cells were centrifuged to assay detergent insolubility, after which proteins were methanol-precipitated from the pellet and supernatant fractions and subjected to proteinase K digestion as described above.
Following digestion, proteins were concentrated by methanol precipitation and analyzed for moPrP by immunoblotting using 3F4 antibody.
Phase Partitioning in Triton X-114
Metabolically labeled cells were solubilized for 20 min at 4 °C in 1% Triton X-114 in phosphate-buffered saline (PBS) containing protease inhibitors; the detergent was diluted from a 12% stock solution that had been precondensed according to Bordier (33). After incubation at 37 °C for 10 min, aqueous and detergent phases were separated by centrifugation. The detergent phase was diluted to the initial volume with PBS, incubated with or without PIPLC for 2 h at 4 °C, and the phase separation repeated. PrP in each phase was immunoprecipitated with antibody 3F4 and separated on SDS-PAGE.
Binding to Phenyl-SepharoseMetabolically labeled cells were lysed and subjected to an initial phase partitioning in Triton X-114 as described above, except that the final detergent concentration was 0.5%. The detergent phase was diluted to the original volume with PBS, incubated with or without PIPLC for 2 h at 4 °C, and then phenyl-Sepharose CL-4B (40 µl of beads, equilibrated in PBS plus 0.5% Triton X-114, per ml of sample) was added for 30 min at 4 °C. The phenyl-Sepharose beads were then collected by centrifugation and bound protein released by boiling in Triton X-100/deoxycholate lysis buffer containing 0.5% SDS. MoPrP in the bound and unbound fractions was analyzed by immunoprecipitation with 3F4 antibody.
Assay of Phospholipase ReleaseFor the experiments shown in Fig. 1, A-B, cells were biotinylated with sulfo-biotin-X-NHS as described by Shyng et al. (30) and were then incubated with PIPLC (1 unit/ml in Opti-MEM) at 4 °C for 2 h. MoPrP in incubation media and cell lysates was then assayed for detergent insolubility or phase partitioning in Triton X-114, as described above. Immunoprecipitates collected using antibody P45-66 were separated by SDS-PAGE, and biotinylated moPrP was visualized by developing blots of the gels with horseradish peroxidase-streptavidin and enhanced chemiluminescence (ECL).
Sucrose Gradient SedimentationLysates of metabolically labeled cells were cleared by centrifugation at 16,000 × g for 2 min and were then loaded on a 10-ml linear gradient of 15-40% sucrose in lysis buffer. After centrifugation at 4 °C for 16 h at 274,000 × g, 1-ml fractions of the gradient as well as the entire pellet were collected, and moPrP was immunoprecipitated with 3F4 antibody.
We have previously shown that moPrP molecules carrying disease-related mutations display several biochemical properties characteristic of PrPSc when expressed in CHO cells (26-28). These properties include (a) detergent insolubility, (b) protease resistance, (c) aberrant membrane attachment, as manifested by retention of the mutant protein on the cell surface after treatment with the enzyme phosphatidylinositol-specific phospholipase C (PIPLC), and (d) hydrophobicity, as revealed by partial retention of the PIPLC-treated protein in the detergent phase of Triton X-114 lysates. Our objective was to use pulse-chase metabolic labeling to analyze the kinetics with which each of these properties was acquired following synthesis of the protein. In this way, we hoped to define a sequence of steps in the transformation of mutant PrPs to a PrPSc-like state.
An important assumption underlying this strategy is that the same population of molecules eventually acquires all four properties. To test this assumption, we assayed the properties in pair-wise fashion using biotinylation or Western blotting to visualize the steady-state population of PrP molecules. For these experiments we utilized CHO cell lines expressing two different moPrP mutants whose human homologues are associated with familial CJD (4): PG11, which contains six additional copies of the N-terminal octapeptide repeat which is found in five copies in the wild-type protein, and E199K which contains a single amino acid substitution.
E199K moPrP was particularly useful for experiments involving PIPLC treatment, since ~50% of this protein is releasable by the phospholipase (compared with <10% for the other mutants), allowing us to analyze both released and non-released fractions (27). We have previously shown that the pool of E199K moPrP that is retained on the cell surface after PIPLC treatment is protease-resistant, whereas the released pool is protease-sensitive (27). To test the correlation between PIPLC releasability and detergent insolubility, we treated surface-biotinylated cells with the phospholipase and then subjected the released and cell-associated fractions to centrifugation at 265,000 × g. We found that the PIPLC-released fraction was found in the supernatant after centrifugation, whereas the retained fraction was primarily in the pellet (Fig. 1A, lanes 5 and 8). To correlate PIPLC releasability and hydrophobicity, we subjected the released and cell-associated fractions of E199K moPrP to Triton X-114 phase partitioning. We found that the released pool partitioned into the aqueous phase, and the cell-associated pool was found in the detergent phase (Fig. 1B, lanes 5 and 8). In contrast to E199K moPrP, wild-type moPrP was completely PIPLC releasable, detergent-soluble, and hydrophilic in Triton X-114 (Fig. 1, A-B, lanes 1-4).
We used PG11 moPrP to test the correlation between detergent insolubility and protease resistance. This mutant protein partitions about equally into pellet and supernatant fractions after centrifugation at 265,000 × g, but it is only the pellet fraction that yields a Mr 27-30 fragment after proteinase K treatment (Fig. 1C, lanes 7 and 8). As expected, wild-type moPrP was found entirely in the supernatant and was protease-sensitive (Fig. 1C, lanes 1-4).
We conclude from these experiments that mutant PrP molecules that are protease-resistant, detergent-insoluble, and hydrophobic are also PIPLC non-releasable. In addition, molecules that are protease-resistant are also detergent-insoluble. Although we have not assayed all four properties simultaneously in the same experiment, our pairwise correlations indicate that at steady-state the same population of mutant moPrP molecules possesses the four biochemical properties of PrPSc.
Intermediate Stages of Aggregation of Mutant PrPThe
centrifugation conditions we have used to assay detergent insolubility
would pellet PrP aggregates having a sedimentation coefficient >40 S.
To more accurately characterize the size distribution of mutant PrP, we
pulse-labeled cells expressing PG11 moPrP and then fractionated
detergent lysates by velocity sedimentation on 15-40% sucrose
gradients. Consistent with earlier results showing that acquisition of
detergent insolubility is primarily a posttranslational process (28),
PG11 moPrP sedimented as a single peak at ~4 S (corresponding to the
size of monomeric protein) immediately following pulse labeling for 20 min (Fig. 2, C-D). After a 2-h chase, the majority of the protein was found in two peaks at 4 S and >16.6 S but a substantial amount was also broadly distributed throughout the
gradient (fractions 5-10). Wild-type moPrP sedimented near 4 S at
both time points (Fig. 2, A-B). The broad size distribution of the PG11 protein at the 2-h chase point indicates that intermediate stages of aggregation of mutant PrP exist and suggests that acquisition of detergent insolubility may be a gradual process in which PrP aggregates grow in size over time. Based on a comparison with molecular
size standards, the smallest aggregates consist of only a few molecules
of PrP, whereas the largest contain over 10.
Detection of Detergent-insoluble, Protease-sensitive PrP
In a
previous study we showed that following pulse labeling PG11 moPrP
becomes maximally detergent-insoluble at 1 h and maximally protease-resistant at 6 h (28). This result suggests that
acquisition of detergent insolubility may precede acquisition of
protease resistance and raises the possibility that a
detergent-insoluble but protease-sensitive form of PrP might be
detectable at early times after pulse labeling. To test this
possibility, we labeled cells expressing PG11 moPrP for 30 min, chased
them for either 0 or 3 h, and then tested the protease resistance
of pellet and supernatant fractions derived from a 265,000 × g centrifugation (Fig. 3). Immediately after
labeling, 33% of the mutant PrP pelleted (lanes 1 and
3), and this amount increased to 65% after a 3-h chase
(lanes 5 and 7). Of note, the insoluble PrP
present immediately after labeling was protease-sensitive, with <1%
of the pelleted protein remaining after proteinase K digestion
(lane 4). In contrast, 15% of the insoluble protein was
protease-resistant following a 3-h chase (lane 8). We
conclude from this result that following synthesis mutant PrP is
initially converted into a form that is detergent-insoluble and
protease-sensitive and only later acquires protease resistance.
Brefeldin A and 18 °C Incubation Inhibit Acquisition of Detergent Insolubility and Protease Resistance
To define the
cellular compartments in which mutant PrPs become detergent-insoluble
and protease-resistant, we utilized two treatments that inhibit the
trafficking of proteins along the secretory pathway. First, we
incubated cells expressing PG11 moPrP with the fungal metabolite
brefeldin A (BFA) which causes fusion of the ER with the
cis/medial-Golgi and thereby inhibits movement of proteins out of these
compartments (34). Second, we incubated cells at 18 °C, a treatment
that blocks movement of proteins from the trans-Golgi network to the
cell surface (35, 36). We found that both of these manipulations caused
a marked inhibition of the acquisition of both detergent insolubility
(Fig. 4) and protease resistance (Fig.
5). The efficacy of BFA and 18 °C incubation in
blocking movement of protein along the biosynthetic pathway was
confirmed by the fact that these treatments inhibited maturation of PrP
oligosaccharide chains (note the lower molecular weight of the two
glycosylated PrP species in Figs. 5, B-C, compared with
Fig. 5A, lanes 1, 3, and 5); in
addition, these treatments prevented delivery of wild-type moPrP to the
cell surface as assayed by PIPLC accessibility (data not shown). We
conclude from our results that transformation of PG11 moPrP to a
detergent-insoluble and protease-resistant form occurs primarily in a
post-Golgi compartment.
We observed that in the presence of BFA a small amount of protease-resistant PrP was detectable after 4 h of chase (Fig. 5, C-D). This result was not likely to be a consequence of escape from the BFA block, since the protease-resistant protein was underglycosylated (23-27 kDa), and was therefore probably derived from molecules that were still trapped in the mixed ER-Golgi compartment (Fig. 5C, lane 6). Similarly, a small percentage of PG11 moPrP became detergent-insoluble after a 4-h chase in the presence of BFA (data not shown). Taken together, these results suggest that detergent insolubility and protease resistance can develop in the ER or Golgi in the presence of BFA, albeit more slowly than under control conditions. Interestingly, no detergent-insoluble or protease-resistant PrP is observed even after prolonged incubation at 18 °C (Fig. 5B, lane 6; and data not shown), possibly because of blockade of a second temperature-sensitive step that is separate from trans-Golgi network-cell surface transport (19).
Identification of an Early Intermediate in the Transformation of Mutant PrPWe sought to identify a step in the conversion of mutant PrP to a PrPSc-like state that preceded the acquisition of detergent insolubility and therefore occurred within 30 min after synthesis of the protein. A clue that such a step might exist was our previous observation that newly synthesized PG11 moPrP molecules are resistant to PIPLC release by the time they reach the cell surface (26), suggesting that an alteration in membrane topology might be an early event in the metabolism of the mutant protein, and one that may occur along the secretory pathway. Since it was technically difficult to test the PIPLC releasability of molecules prior to their arrival at the plasma membrane, we chose instead to assay the hydrophobicity of the protein after treatment with PIPLC, a characteristic that was likely to correlate with aberrant membrane attachment. We had previously shown that, after treatment with PIPLC, surface-biotinylated PG11 moPrP is partially retained in the detergent phase following Triton X-114 phase partitioning, whereas wild-type PrP is shifted almost entirely into the aqueous phase (26).
We asked when after pulse labeling the PG11 molecule first displays
this aberrant behavior in Triton X-114 phase partitioning (Fig.
6, lanes 1-8). We found that both wild-type
and PG11 moPrP partitioned into the detergent phase prior to PIPLC
treatment, a behavior that is expected because of the fatty acyl or
alkyl chains present on the glycosylphosphatidylinositol (GPI) anchors of both proteins. After PIPLC treatment, wild-type moPrP was shifted entirely into the aqueous phase because of cleavage of the anchor. In
contrast, approximately half of the PG11 moPrP protein was retained in
the detergent phase following phospholipase treatment. This result may
be attributable to failure of PIPLC to remove the GPI anchor from the
PG11 molecule or to an intrinsic hydrophobicity of the polypeptide
chain after the anchor has been removed (26). Significantly, retention
in the detergent phase was seen after a 20-min pulse labeling period
with no chase, arguing that whatever molecular change was responsible
for this aberrant behavior in Triton X-114 occurred very early after
synthesis of PG11 PrP molecules.
We used binding to phenyl-Sepharose as a second method to assay the
hydrophobicity of PG11 molecules after PIPLC treatment (Fig.
7, lanes 1-4). This resin binds hydrophobic
molecules, and the distribution of a protein between bound and unbound
fractions is a measure of its hydrophobic character (37, 38). As
expected because of the presence of the GPI anchor, both wild-type and PG11 moPrP were concentrated in the bound fraction prior to PIPLC treatment. Incubation with PIPLC shifted all of the wild-type protein
into the unbound fraction, whereas only half of the PG11 protein was
shifted. Retention of PG11 moPrP in the bound fraction was observed
immediately following the 30-min labeling period in the absence of a
chase. These results are analogous to those obtained using partitioning
in Triton X-114 to assay hydrophobicity.
We obtained similar results when cells expressing PG11 moPrP were pulse-labeled for as little as 10 min prior to Triton X-114 phase partitioning or phenyl-Sepharose fractionation (data not shown).
Brefeldin A Does Not Affect Triton X-114 Phase Partitioning or Phenyl-Sepharose BindingOur results suggested that the partial hydrophobicity of PG11 PrP following PIPLC treatment was a feature that developed during or very soon after synthesis of the polypeptide chain, at a time when the protein is likely to reside in the ER or Golgi. Consistent with this hypothesis, we found that BFA had no effect on the behavior of PG11 moPrP as assessed by either Triton X-114 phase partitioning (Fig. 6, lanes 9-16) or binding to phenyl-Sepharose (Fig. 7, lanes 5-8); in both assays, about half of the molecules remained hydrophobic after PIPLC treatment whether or not BFA was present. These results indicate that whatever molecular change alters the response of the PG11 molecule to PIPLC in the two assays is likely to develop within the ER or cis/medial-Golgi.
We have shown previously that moPrP molecules carrying
disease-related mutations display a number of biochemical markers
characteristic of PrPSc when expressed in CHO cells
(26-28). Our objective in this study was to examine the time course
over which each of these operational properties is acquired after
synthesis of the protein, with a view to defining intermediate stages
in the acquisition of the PrPSc-like state. We have assayed
three different biochemical hallmarks of the scrapie isoform and found
that although all of them are present in mutant PrP molecules at steady
state, each property develops with different kinetics in pulse-chase
experiments. In addition, we have used inhibitors of protein
trafficking to help pinpoint the cellular compartments where these
biochemical transformations take place. Our results suggest the model
shown in Fig. 8. We propose that mutant PrPs are
initially synthesized in the PrPC state and then acquire
the characteristics of PrPSc in a stepwise fashion during
passage through various cellular compartments.
The earliest biochemical change we could detect in mutant PrP was a partial resistance to the effect of PIPLC in rendering the protein hydrophilic. About half of the mutant protein remained hydrophobic after treatment with the phospholipase, as assessed by partitioning in Triton X-114 or binding to phenyl-Sepharose. In contrast, essentially all of the wild-type protein was converted to a hydrophilic form. Our initial investigations suggested that this abnormal behavior of PG11 moPrP was due to an intrinsic hydrophobicity of the mutant polypeptide chain, since we showed that PIPLC had released 3H-fatty acid label from the GPI anchor of many of the molecules (26). However, more recent studies have suggested that an additional explanation may be that the anchors of some of the PG11 molecules are inaccessible to phospholipase cleavage unless the protein is denatured.3 Whatever the relative importance of the two mechanisms, however, the unusual behavior of mutant PrP following PIPLC treatment was already observable in molecules that had been pulse-labeled for only 10 min and was not affected by treatment of cells with BFA. These observations suggest that an initial alteration in the mutant PrP molecule occurs in the ER, during or very soon after translation of the polypeptide chain. We cannot, however, rule out the possibility that this change takes place in the cis or medial compartments of the Golgi, which become fused with the ER after BFA treatment (34). We think it is probable that abnormal behavior in the Triton X-114 and phenyl-Sepharose assays correlates with inefficient release of mutant PrPs from the cell membrane by PIPLC, since both properties develop prior to arrival of the protein at the cell surface, and since both are likely to reflect an alteration in membrane association (26).
The second step we have identified is acquisition of detergent insolubility. Some detergent insolubility is detectable at the end of a 30-min labeling period (Fig. 3), but this property is not maximal until 1 h of chase (28), arguing that it occurs after the alteration in hydrophobicity properties. Consistent with this proposal, development of detergent insolubility is inhibited by BFA and incubation at 18 °C, suggesting that it occurs in a post-Golgi location. The relevant sites remain to be identified, but they could include the plasma membrane, caveolae-like microdomains (39, 40), or endosomes. Detergent insolubility presumably reflects aggregation of PrP molecules, and by sucrose gradient fractionation we were able to detect aggregates ranging in size from 4 S (monomeric) to >16.6 S (>10 PrP molecules). Although we have not directly proven that smaller aggregates are converted to larger aggregates, we speculate that PrP oligomers exist in intact cells and grow in size over time. Whether this process has the properties of a nucleated polymerization, as proposed by some investigators (12, 14-16), remains to be proven.
The third step in the transformation of mutant PrP is acquisition of protease resistance. This property is not maximal until several hours after labeling (28), and its development is blocked by BFA treatment and 18 °C incubation (Fig. 5), consistent with it happening in a post-Golgi location. We have shown that mutant PrP is detergent-insoluble and protease-sensitive at early times after pulse labeling but then becomes protease-resistant with subsequent chase. This result directly demonstrates that acquisition of detergent insolubility and protease resistance are temporally distinct steps connected by an intermediate state. It is easy to imagine, however, that the two steps might be related if, for example, PrP aggregates became protease-resistant upon reaching a certain minimum size. Gabizon et al. (41) have detected a form of wild-type PrP that is detergent-insoluble but protease-sensitive in the brains of heterozygous patients carrying an E200K mutation, although it was not possible in their experiments to determine by metabolic labeling whether this was a true intermediate that could be converted into a protease-resistant form. Priola et al. (42) have described a 60-kDa covalently linked dimer of PrP that is aggregated but protease-sensitive, but we have not observed this species in our cells.
Although we have shown that mutant PrPs synthesized in CHO cells display all the biochemical hallmarks of PrPSc, we are still in the process of testing whether the proteins are infectious in animal bioassays. If the proteins should turn out to be infectious, it will be important to determine at what point in the pathway shown in Fig. 8 this critical attribute is acquired. Infectivity could develop simultaneously with the earliest biochemical changes that we hypothesize take place in the ER, or it may represent an additional step that depends on PrP aggregates growing beyond a certain size, or attaining a minimum level of protease resistance. Even if the mutant PrPs from CHO cells should turn out to lack infectivity, it seems likely that the steps presented in Fig. 8 represent necessary prerequisites for acquisition of this property.
Our data provide the first evidence that conversion to the PrPSc state may begin at an early point in the biosynthetic pathway, perhaps even simultaneously with translation of the PrP polypeptide chain in the ER. Previous studies have emphasized events occurring subsequent to arrival of PrP molecules at the cell surface (18, 19). A role for the ER in transformation of mutant PrPs is appealing from a theoretical standpoint because of the well known role of this organelle in protein folding and glycosylation. Recent evidence suggests that prion strains are distinguished by differences in utilization of the two consensus sites for N-linked glycosylation (43-45), and it is possible that these differences arise in the ER as a result of variations in the efficiency with which oligosaccharyl transferase transfers the dolichol-linked precursor to nascent PrP chains. It is also tempting to speculate that chaperone molecules in the ER, which catalyze the folding of newly synthesized polypeptide chains (22), play an important role in the conformational changes that are thought to underlie generation of PrPSc. Mutant glycoproteins associated with several other human genetic diseases are known to associate abnormally with ER chaperones (46), and it will be interesting to see if the same is true for mutant PrPs. Several cytoplasmic heat-shock proteins that function as molecular chaperones have been implicated in prion phenomena in scrapie-infected cells (47) and in yeast (48), although there has not been any experimental investigation of the role of ER chaperones. Recent genetic experiments have also been interpreted to suggest the involvement of accessory proteins, some of which may be molecular chaperones, in the generation of the PrPSc (23).
Our results invite speculation about the relationship between the
operational biochemical parameters we have assayed and the fundamental
conformational change that is thought to underlie conversion of
PrPC into PrPSc. Spectroscopic data indicate
that PrPSc has a higher content of -sheets and a lower
content of
-helices than PrPC, and it has been proposed
that conversion of
-helices into
-sheets is responsible for
generation of the scrapie isoform (5-8). We favor the possibility that
this conformational transition happens either during or soon after
synthesis of mutant PrP molecules in the ER and that a direct
manifestation of this change is the aberrant response of the protein to
PIPLC in the Triton X-114 and phenyl-Sepharose assays. In this view,
detergent insolubility and protease resistance are secondary properties
that develop only some time after the initial molecular conversion. On
the other hand, we cannot rule out the possibility that the altered behavior of the protein in the two hydrophobicity assays is unrelated to the fundamental change in PrP conformation, which might occur later
and be directly responsible for development of insolubility and
protease resistance. It is also conceivable that the conformational change occurs in a gradual rather than an all-or-none fashion and that
each of the three biochemical transitions we have observed is related
to an incremental change in folding of the PrP molecule.
We have assumed that each of the biochemical transformations shown in Fig. 8 depends on transit of the mutant PrP molecule through a specific cellular compartment. A particular subcellular location may provide a favorable environment for a given transition because of the presence of cofactor molecules, for example chaperones in the ER, or because of physical conditions, like the acidic milieu of endocytic organelles. However, it is also likely that mutant PrPs have some intrinsic tendency to acquire PrPSc-like properties independent of the cellular compartment where they reside. We observed that in the presence of BFA, PG11 moPrP started to become detergent-insoluble and protease-resistant after a prolonged period of chase (Fig. 5C lane 6, and data not shown). This result indicates that, although these two properties are normally acquired in a post-Golgi location, they can be forced by pharmacological manipulation to develop earlier in the biosynthetic pathway. In addition, we have found that a detergent-insoluble form of PG11 moPrP can be generated by incubating detergent lysates of pulse-labeled cells at 37 °C for many hours, a situation in which all cellular organelles are disrupted.2 In a similar vein, it has been demonstrated recently that protease-resistant PrP can be produced in vitro using purified proteins or synthetic peptides, although the efficiency of this process is low (12, 13). Taken together, these results argue that the generation of PrPSc involves intrinsic structural features of the PrP molecule itself but that the efficiency of the process is greatly promoted by cell- and organelle-specific factors.
A number of observations suggest that the scheme we have defined here for transformation of mutant PrPs also describes the conversion of wild-type PrP into PrPSc in cells infected with exogenous prions. In scrapie-infected neuroblastoma and immortalized hamster brain cells, BFA treatment and 18 °C incubation also block the appearance of protease-resistant PrP (19, 49). A similar effect is seen when cells are treated with PIPLC and proteases, which presumably remove or inactivate the PrPSc precursor on the cell surface (18, 19). These results have led to the proposal that PrPSc is generated either on the cell surface, perhaps in cholesterol-rich microdomains (20, 21), or along the endocytic pathway. In light of the results reported here, we would propose that acquisition of detergent insolubility and protease resistance occur in these post-Golgi locations but that there may be an earlier step that takes place in infected cells prior to arrival of PrP at the cell surface. By analogy to mutant PrP, this step might take place in the ER and be detectable experimentally by testing the ability of PIPLC to convert the PrPSc precursor to a hydrophilic form and to completely release it from the cell membrane. In fact, we have already shown that at steady state PrPSc in infected neuroblastoma cells is retained on the cell surface after PIPLC treatment (27). It will be important now to carefully compare the intermediate steps underlying PrPSc production in infected cells with those operative in cells expressing mutant PrPs to define differences and similarities between the infectious and inherited manifestations of prion diseases.
We thank Rick Kascsak for antibodies as well as Cy Pauly and Maurine Linder for critical evaluation of the manuscript.