1 Institute for Neurodegenerative Diseases, University of California, 513 Parnassus Ave, San Francisco, CA 94143, USA
2 Department of Neurology, University of California, 513 Parnassus Ave, San Francisco, CA 94143, USA
3 Department of Pathology, University of California, 513 Parnassus Ave, San Francisco, CA 94143, USA
4 Department of Biochemistry and Biophysics, University of California, 513 Parnassus Ave, San Francisco, CA 94143, USA
Correspondence
Stanley B. Prusiner
stanley{at}ind.ucsf.edu
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ABSTRACT |
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Supplementary material is available in JGV Online.
Present address: Neurochem Inc., 275 boul. Armand-Frappier, Laval, Quebec, Canada H7V 4A7.
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INTRODUCTION |
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Prions in both mammals and yeast multiply by inducing the precursor protein to adopt the conformation of the protein in the prion state. In fungi, polymerizing the prion precursor proteins into amyloid fibrils is sufficient to create infectious particles (Maddelein et al., 2002; Sparrer et al., 2000
). In mammals, the task seemed more complex until recently (Legname et al., 2004
).
In attempting to produce mammalian prions de novo, we initially employed transgenic (Tg) mice expressing high levels of MoPrP(P101L), which harbours the analogous mutation causing GerstmannSträusslerScheinker syndrome in humans. These mice developed neurodegeneration spontaneously (Hsiao et al., 1990) and brain extracts from these mice transmitted prion disease to Tg mice expressing low levels of MoPrP(P101L), designated Tg196 mice (Hsiao et al., 1994
; Telling et al., 1996
). Such experiments were plagued by the inability to demonstrate protease-resistant (r)PrPSc(P101L) in the brains of the spontaneously ill Tg mice, as well as of the inoculated Tg196 mice. Subsequently, we used the P101L mutation to guide the folding of a 55 aa peptide composed of MoPrP residues 89143 into a
-rich conformation that polymerized into fibrils. The fibrils produced neurological dysfunction
1 year after inoculation into Tg196 mice, while the non-
-rich form of this peptide did not (Kaneko et al., 2000
). Brain extracts prepared from ill Tg196 mice serially transmitted disease to Tg196 recipients with a similar incubation time, but did not cause disease in wild-type (wt) mice (Tremblay et al., 2004
).
Impressed by the amyloid deposition in the brains of Tg196 mice inoculated with -rich fibrils (Tremblay et al., 2004
), we investigated larger, recombinant (rec)PrPs polymerized into amyloids, which represent a subset of
-rich structures. Using recMoPrP(89230) produced in Escherichia coli to form amyloid fibrils, we inoculated the fibrils intracerebrally into Tg mice expressing MoPrP(89231) (Supattapone et al., 2001a
). These Tg mice developed neurological dysfunction between 380 and 660 days after inoculation, and exhibited rPrPSc and spongiform degeneration in their brains, homogenates of which transmitted disease to wt and Tg mice expressing full-length PrP (Legname et al., 2004
).
The production of both wt and mutant synthetic prions established that PrPSc is the sole component of the mammalian prion. While Tg mouse studies have demonstrated that the rate of PrPSc accumulation in brain is directly proportional to the level of PrPC expression (Prusiner et al., 1990), little is known about the factors that govern the rates of PrPSc formation and clearance. The availability of bigenic mice, designated Tg(tTA : PrP+/0)3, in which PrPC expression can be reversibly regulated (Tremblay et al., 1998
), offers an opportunity to investigate the formation and clearance of PrPSc in vivo. Tg(tTA : PrP+/0)3 mice were inoculated with RML prions and developed neurological dysfunction
150 days later. When PrPC expression was suppressed by
95 % by administration of doxycycline, the onset of neurological deficits was delayed to
430 days (Tremblay et al., 1998
). Using these bigenic mice, we measured the clearance as well as the rates of accumulation of both rPrPSc and protease-sensitive (s)PrPSc in brain. In the brains of these prion-infected mice, the half-life (t1/2) for clearance of PrPSc was
1·5 days, in good agreement with studies of scrapie-infected neuroblastoma (ScN2a) cells (Peretz et al., 2001b
).
The capability of the brain to clear prions raises the possibility that PrPSc is normally made at low levels and continually cleared. The steady-state level of PrPSc must be below 105 molecules per ml of 10 % (w/v) brain homogenate (BH) since homogenates prepared from uninfected animals do not transmit disease to inoculated recipients (Prusiner et al., 1983). The hypothesis that PrPSc might be present at low levels in normal animals raises the question as to whether PrPSc has an as-yet-unidentified metabolic function. Moreover, this conjecture suggests that prion diseases may arise from dysregulation in the concentration of PrPSc rather than the formation of a new conformer.
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METHODS |
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Prion inoculum.
Anaesthetized animals were inoculated at 8 weeks (±4 days) of age with 20 µl RML prions (1 % BH in PBS) and observed for signs of neurological dysfunction three times a week as described previously (Carlson et al., 1994; Scott et al., 1989
). Mice with fully developed clinical symptoms were euthanized and their brains harvested for further studies.
Administration of doxycycline and care of laboratory animals.
For breeding, animals were kept on 0·02 mg doxycycline ml1 in their drinking water, which was ceased at 3 weeks of age to allow high levels of PrP expression (Tremblay et al., 1998). After animals were inoculated intracerebrally with RML prions, doxycycline administration was initiated at different time points post-inoculation to suppress PrPC expression. In dose-dependency experiments, doxycycline at 0·002, 0·02, 0·2 or 2 mg ml1 was added to the drinking water with 5 % sucrose to mask the bitter taste. In clearance experiments, an initial single dose of 25 mg doxycycline kg1 was administered intraperitoneally, followed by 2 mg doxycycline ml1 in the drinking water.
Treatment of BHs.
For biochemical analysis only, 10 % BH samples were prepared in Ca2+/Mg2+-free PBS by homogenization (three times for 15 s each) with a PowerGen 125 homogenizer (Fisher Scientific). Homogenates were clarified by centrifugation at 500 g for 5 min on a tabletop centrifuge. Supernatants were subjected either to sodium phosphotungstate (PTA) precipitation or to proteinase K (PK; Gibco-BRL) treatment followed by PTA precipitation (Safar et al., 1998). For PK digestion, 5 % BH samples were incubated with 25 µg PK ml1 for 1 h at 37 °C. PK activity was blocked using a protease inhibitor (PI) cocktail, composed of 2 µg aprotinin and leupeptin (A/L) ml1 and 0·2 mM PMSF. Samples were diluted with one volume of 4 % Sarkosyl/PBS and analysed by Western blotting or by conformation-dependent immunoassay (CDI) (see below).
For PK digestion followed by PTA precipitation, 10 % BH was treated with PK and the reaction was blocked using the PI cocktail. The samples were diluted with one volume of 4 % Sarkosyl/PBS and then PTA precipitated. For PTA precipitation, 1 ml aliquots of the samples were adjusted to 0·31 % PTA, 2·6 mM MgCl2, incubated at 37 °C for 116 h and centrifuged at 16 000 g for 30 min. Pellets were resuspended in PBS with PI and 0·2 % Sarkosyl and immediately analysed by Western blotting or CDI (see below).
Western blotting.
Volumes (500 µl) of 5 % (w/v) homogenates were prepared in PBS containing 2 % (w/v) Sarkosyl and the protein concentration was measured using the bicinchoninic acid assay (Pierce). For digested samples, aliquots were treated with 20 µg PK ml1 at a ratio of 1 : 50 [PK : protein (w/w)] for 1 h at 37 °C. Digestion was stopped with 5 mM PMSF. PK-treated samples were mixed with an equal volume of SDS loading buffer and boiled for 5 min; 30 µl samples were analysed by 12 % SDS-PAGE using precast gels (Bio-Rad). For undigested samples, aliquots were mixed with an equal volume of SDS loading buffer and boiled for 5 min; 10 µl samples were analysed by 12 % SDS-PAGE using precast gels. Western blots were developed with 1 µg recFab D13 ml1 and peroxidase-labelled anti-Fab secondary antibody, and detected using the enhanced chemiluminescence system as described previously (Peretz et al., 2001a).
Detection of sPrPSc and rPrPSc by CDI.
Samples were processed for CDI using a time-resolved fluorescence (TRF) method as described previously (Safar et al., 1998). Samples were split into two aliquots; one was kept untreated (native) and the other (denatured) was treated with one volume of 8 M guanidinium hydrochloride and heated at 80 °C for 5 min. Both aliquots were diluted 20-fold with water containing 0·5 mM PMSF and 2 µg A/L ml1. Samples were loaded in triplicate on 96-well polystyrene microplates (OptiPlate HTRF-96; Packard) that had been pre-coated with recFab D18 (Peretz et al., 1997
) for the sandwich-formatted CDI or pre-activated with glutaraldehyde (0·2 % in PBS, pH 7·4; 2 h) for direct CDI. The plates were incubated for 2 h at room temperature and blocked overnight at 4 °C with Tris-buffered saline [TBS; 20 mM Tris/HCl pH 7·5; 1 % BSA; 6 % (w/v) sorbitol]. The next day,
-PrP europium-labelled, chimeric humanmouse (HuM) recFab D13 (Peretz et al., 1997
) at a concentration of 0·55 µg ml1 was added to the plates and incubated for 2 h at room temperature. The (DN) difference, which is the binding of antibody to native and denatured samples, is directly proportional to the concentration of PrPSc. The concentration may be determined by a mathematical model that was developed previously to calculate the amount of
-sheet in PrP (Safar et al., 1998
). The concentration of PrP in the samples was calculated from calibration curves present on each plate and generated with
-rich recMoPrP(89231) (Mehlhorn et al., 1996
) serially diluted into BH of Prnp0/0 transgenic mice (Büeler et al., 1992
). The concentration of sPrPSc was calculated according to the formula: [sPrPSc]=[PrPSc][rPrPSc], in which [PrPSc] is the concentration of PrPSc before PK treatment and [rPrPSc] is the concentration of the protease-resistant fraction after PK treatment.
Neuropathology.
Brains were rapidly removed from animals and either perfusion fixed in 10 % buffered formalin or snap frozen. Immunohistochemical localization of rPrPSc was accomplished on formalin-fixed, paraffin-embedded tissue sections by the formic acid-hydrated autoclaving method (Muramoto et al., 1997) and on cryostat sections by the histoblot procedure (Taraboulos et al., 1992a
). HuM D13 and HuM D18, which recognize MoPrP(97106) and MoPrP(132156), respectively (Peretz et al., 1997
), were used as primary antibodies.
Cryostat sections were pressed on to wet nitrocellulose paper and allowed to air dry. Sections were exposed to 200 µg PK ml1 for 1 h at 37 °C to eliminate PrPC, followed by denaturation of the remaining rPrPSc with 3 M guanidine isothiocyanate for 10 min at room temperature. Histoblots were then incubated overnight at 4 °C with D13 or D18 diluted 1 : 100. Following rinsing, histoblots were incubated with the secondary antibody, alkaline phosphatase-conjugated anti-human IgG (Promega), for 1 h at room temperature and colour was developed with NBT/BCIP (nitro blue tetrazolium/5-bromo-4-chloro-3-indolyl phosphate) substrate (Promega) after additional rinses.
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RESULTS |
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Kinetics of PrPSc accumulation in the brains of Tg(tTA : PrP+/0) mice
To study the clearance of PrPSc from the brains of Tg(tTA : PrP+/0)3 mice, we first determined the kinetics of PrPSc accumulation after intracerebral inoculation with RML prions. Tg(tTA : PrP+/0)3 mice were sacrificed at various intervals after inoculation and PrPSc levels determined by CDI (Supplementary Fig. S1a). PrPSc levels peaked at 98 days post-inoculation and gradually decreased over the next 40 days. By 142 days post-inoculation, the PrPSc level in the brains of these asymptomatic mice was 80 % of the level found at 98 days. From this point, the mice remained well for another week until neurological dysfunction ensued (Fig. 1
). In contrast to the biphasic accumulation of PrPSc, the accumulation of rPrPSc was monophasic and exponential with a maximum level found at 142 days, approximately 1 week before the onset of neurological symptoms (Supplementary Fig. S1b). We did not measure the levels of rPrPSc during the clinical phase of disease; some investigators have argued that prion titres rise during this period, while others have found no evidence to support such a contention (Brown et al., 1982
; Oesch et al., 1985
).
Doxycycline modulates PrPSc levels in Tg(tTA : PrP+/0) mice
At five different times after inoculation of RML prions, doxycycline was administered to Tg(tTA : PrP+/0)3 mice. Starting doxycycline at 98 or 126 days after inoculation demonstrated that readily measurable reductions in PrPSc could be achieved (Supplementary Fig. S2a). Similarly, reductions in rPrPSc could be achieved by doxycycline administration beginning at both 98 and 126 days after inoculation, as well as at 84 days (Supplementary Fig. S2b). Administration of doxycycline at the midpoint of the incubation time, at 70 and 77 days after inoculation, did not create conditions for measuring decrements in the level of either PrPSc or rPrPSc, and the mice were asymptomatic at 290 days post-inoculation. For comparison, mice expressing PrPC inoculated with RML prions developed disease at 150 days (Fig. 1
).
Clearance of sPrPSc and rPrPSc
Based on the clearance curves in Supplementary Fig. S2, we administered doxycycline to Tg(tTA : PrP+/0)3 mice beginning 98 days after inoculation with RML prions. At six different time points, we sacrificed Tg(tTA : PrP+/0)3 mice and measured PrPC and PrPSc levels in their brains.
PrPC levels in the brains of uninoculated Tg(tTA : PrP+/0)3 mice declined to 40 % of the initial level within 1 day of beginning doxycycline treatment and to
5 % after 7 days (Figs 2 and 3a
). This low level of PrPC persisted for the remainder of the 56 day period. From CDI measurements (Fig. 2
) and densitometric scans of Western blots (Fig. 3a
) of PrPC levels in the brains of uninfected Tg(tTA : PrP+/0)3 mice, we estimated that the apparent t1/2 of PrPC in the brains of these mice was
18 h.
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Suppression of PrPC results in clearing of rPrPSc from grey matter
To examine the impact of PrPC suppression on the distribution of rPrPSc in the brain, Tg(tTA : PrP+/0)3 mice were inoculated with RML prions, administered doxycycline at 98 days post-inoculation and sacrificed 58 days later. Animals on doxycycline were well when sacrificed, in contrast to inoculated Tg(tTA : PrP+/0)3 mice not receiving doxycycline, which exhibited neurological dysfunction at 156 days post-inoculation. The ill animals exhibited rPrPSc accumulation throughout the brain, with the most intense immunostaining in the white matter tracts of the thalamus (Fig. 5a) and white matter of the corpus callosum and cerebellum (Fig. 5b
). rPrPSc was also found in grey matter, where immunostaining resulted in a low-intensity signal that enabled the neocortex, hippocampus and cerebellar cortex to be delineated on the histoblots. In contrast, the mice receiving doxycycline beginning at 98 days post-inoculation and sacrificed at 156 days post-inoculation exhibited little or no rPrPSc in grey matter (Fig. 5c and d
). Residual rPrPSc was found almost exclusively in white matter, particularly in the corpus callosum (Fig. 5c
). In the cerebellum, rPrPSc immunostaining was seen primarily at the interface of the white matter and cerebellar cortex (Fig. 5d
).
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DISCUSSION |
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Inducible transgenes and PrPSc clearance from the brain
Using Tg(tTA : PrP+/0)3 mice, we demonstrated that PrPSc can be cleared from the brain following doxycycline suppression of PrPC expression between 84 and 126 days after inoculation (Supplementary Fig. S2b). Our studies demonstrate that, in brain, PrPSc is not only being formed but is also being removed. The t1/2 value of clearance of both sPrPSc and rPrPSc was 1·5 days (Table 2
). That both sPrPSc and rPrPSc have similar t1/2 values was unexpected, since rPrPSc is resistant to proteolytic degradation and thus would be expected to have a longer half-life than sPrPSc. It is possible that the actual t1/2 might be shorter due to the lag phase necessary for doxycycline to reach equilibrium in the brain and for complete inhibition of PrP transgene transcription. The lag phase for this inducible transgene system in bigenic mice has been estimated to be <1 h (Gossen & Bujard, 2002
). However, the contribution of continuing PrPSc conversion from residual PrPC expression, which remains at
5 % of the level in Tg(tTA : PrP+/0)3 mice, might increase the t1/2 of PrPSc. We do not believe that this low level of nascent PrPSc formation would substantially change the t1/2 for PrPSc.
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Clearance of PrP isoforms from cultured cells
Anti-PrP antibodies have been used to determine the apparent t1/2 for PrPSc in ScN2a cells (Peretz et al., 2001b). A t1/2 value of
30 h was calculated for the clearance of PrPSc in ScN2a cells (Table 2
), which is similar to the 36 h calculated for PrPSc in brain. It is noteworthy that scrapie-infected cell cultures probably represent a mixture of infected cells and uninfected revertants. Some evidence supports the proposition that infected cells in culture eventually die when their PrPSc levels rise above a threshold level (Schätzl et al., 1997
; Tatzelt et al., 1996
). Whether the t1/2 value of 30 h in ScN2a cells overestimates the actual t1/2 due to the unknown rate of inhibition of PrPSc formation in cells continuously expressing PrPC remains to be established (Peretz et al., 2001b
; Supattapone et al., 2001b
).
Most studies of PrPC turnover and cellular metabolism have relied on cultured cells (Borchelt et al., 1990; Caughey et al., 1989
; Drisaldi et al., 2003
; Gilch et al., 2001
), in which glycosylated and glycolipidated PrPC appears on the cell surface. Whether PrPC reaches the plasma membrane or axon terminal in vivo remains to be determined (Borchelt et al., 1994
). The deduced t1/2 of PrPC derived from experiments using ScN2a cells is relatively short, ranging from 2·6 to 7 h, depending on the experimental design and PrP sequence (Borchelt et al., 1990
; Caughey et al., 1989
; Drisaldi et al., 2003
; Nunziante et al., 2003
). In contrast, the apparent t1/2 of PrPC expressed in mouse brain presented here was
18 h (Table 2
) (Borchelt et al., 1994
; Daude et al., 1997
; Peters et al., 2003
).
The t1/2 value for PrPC represents the mean of different turnover times in diverse cell types in many different brain regions. Some additional factors include the turnover of PrPC in different subcellular compartments of mature neurons, such as PrPC that migrates by fast axonal transport to the nerve terminals, PrPC recently documented in cytosol and PrPC internalized in endosomal compartments via caveolae (Borchelt et al., 1994; Mironov et al., 2003
; Peters et al., 2003
). Although PrP mRNA and PrPC have consistently short t1/2 values in ScN2a cells, it is likely that PrP mRNA in the brains of Tg mice has an extended t1/2, which contributes to the extended t1/2 of PrPC observed in the brain (Borchelt et al., 1990
; Muller et al., 1997
). Moreover, t1/2 measurements of PrPC in ScN2a cells under non-differentiating conditions do not reflect the expected complexity of mRNA and PrPC turnover in a variety of fully differentiated neurons and glial cells in the mature mouse brain in our experiments (Borchelt et al., 1990
; Caughey et al., 1989
; Drisaldi et al., 2003
; Gilch et al., 2001
). The bigenic mouse system described in this paper now permits us to determine PrP mRNA and PrPC turnover at a cellular level in the brain.
Factors modulating PrPSc levels
The findings reported here and previously demonstrate that PrPSc levels in brain are determined by both the rates of formation and clearance. In earlier studies, we demonstrated that PrPC expression is directly proportional to the rate of PrPSc formation (Prusiner et al., 1990), but inversely related to the length of the incubation time: the greater the level of PrPC expression, the greater the rate of PrPSc formation and the shorter the incubation time.
In general, PrPSc formation as reflected in shorter incubation times occurs when the sequences of PrPC and PrPSc are the same (Prusiner et al., 1990; Scott et al., 1989
). As an exception, a particular strain of prion may preferentially interact with PrPC of another sequence. For example, variant CJD prions composed of HuPrPSc are more readily transmitted to Tg(BoPrP) mice expressing bovine PrP than to Tg(MHu2M) mice expressing chimeric mousehuman PrP (Korth et al., 2003
; Scott et al., 1999
). This contrasts with sporadic CJD, familial CJD(E200K) and iatrogenic CJD prions, all of which more readily transmit to Tg(MHu2M) mice than to Tg(BoPrP) mice (Scott et al., 2005
; Telling et al., 1995
). These findings argue that the tertiary structure of PrPSc reflecting the particular prion strain governs the interaction with PrPC during prion replication.
Whether strains of prions are cleared or formed at different rates remains to be established. In one example, mice expressing MoPrP-A or MoPrP-B, which differ at positions 108 and 109, replicated prions at different rates (Westaway et al., 1987).
Mechanism and kinetics of PrPSc clearance from the brain
The mechanism of PrPSc clearance from the brain is unknown. From cultured cell studies, it seems likely that PrPSc is hydrolysed in acidic endosomes as well as lysosomes (Caughey et al., 1990; Taraboulos et al., 1992b
). The clearance of PrPSc was accelerated in ScN2a cells by branched polyamines, presumably by diminishing the resistance of rPrPSc to proteolysis at acidic pH (Supattapone et al., 1999
, 2001b
).
In Tg(tTA : PrP) mice given 2 mg doxycycline ml1 in the drinking water, PrPC expression was reduced by 95 %. The residual PrPC was sufficient to support PrPSc replication, albeit at a slow rate. These bigenic mice eventually developed central nervous system degeneration at
430 days after inoculation (Table 1
). The slow accumulation of PrPSc is reflected in studies of t1/2 for clearance measured both by CDI and Western blot, in which PrPSc rapidly declined after administration of doxycycline and eventually began to accumulate again (Figs 3c and 4a
). The extremely rapid response of Tg(tTA : PrP) mice to oral doxycycline in suppressing PrP mRNA transcription followed by PrPSc degradation suggests that the t1/2 measurements are likely to reflect the actual rates of PrPSc clearance.
Therapeutics for prion disease
The data presented here demonstrate that PrPSc can be cleared from the brain. This is encouraging since it argues that drugs that abolish PrPSc formation or enhance clearance can rid the brain of prions. However, the efficacy of such drugs will depend on how well they accomplish either task: PrPSc formation must be abolished completely or clearance enhanced to remove all PrPSc.
As in earlier studies (Brandner et al., 1996), our data indicate a critical role of PrPC in neurodegeneration caused by PrPSc. We found that residual rPrPSc persisting in some brain areas after suppression of PrPC expression produced few or no signs of neurodegeneration in surrounding cells (Fig. 6
). In the presence of PrPC, a severe loss of pyramidal neurons was observed in the hippocampal CA1 region; moreover, the remaining neurons were shrunken, suggesting that they were also undergoing degeneration. In contrast, no obvious nerve cell loss was detectable in animals in which PrPC expression was suppressed. Therefore, the structural transition that PrPC undergoes should prove to be an effective drug target.
Does PrPSc have a cellular function?
Studies demonstrating the clearance of PrPSc raise the possibility that PrPSc is normally made at low levels and continually removed. Such a proposal posits that PrPSc may have an as-yet-unidentified function and that prion diseases are disorders of PrPSc metabolism.
If we assume that PrPSc is formed in normal cells, then we would argue that prion diseases arise from the dysregulation of PrPSc metabolism. Thus, PrPSc begins to accumulate when the rate of formation exceeds the rate of clearance. As the net accumulation of PrPSc continues to increase, a point is reached when a cell can no longer tolerate the level of PrPSc and it begins to malfunction. Such a scenario seems particularly appealing in the spontaneous and inherited forms of prion disease.
Explaining how the seemingly wide variety of PrPSc conformations, each of which represents a different strain, might participate in regulating the metabolism of normal cells poses a conundrum. However, determining whether prion strains with different incubation times show different rates of PrPSc clearance might prove informative. Defining the rates of both formation and clearance for different prion strains would seem to be a fundamental issue in prion biology, which is now amenable to investigation using the bigenic mouse system described here.
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ACKNOWLEDGEMENTS |
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Received 4 February 2005;
accepted 19 May 2005.
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