From the Laboratory of Persistent Viral Diseases,
Rocky Mountain Laboratories, NIAID, National Institutes of Health,
Hamilton, Montana 59840 and the § Chemistry Department, The
University of Montana, Missoula, Montana 59812
Received for publication, November 13, 2002, and in revised form, January 24, 2003
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ABSTRACT |
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A key event in the pathogenesis of transmissible
spongiform encephalopathies is the conversion of PrP-sen to PrP-res.
Morrissey and Shakhnovich (Morrissey, M. P., and Shakhnovich,
E. I. (1999) Proc. Natl. Acad. Sci. U. S. A. 96, 11293-11298) proposed that the conversion mechanism involves critical
interactions at helix 1 (residues 144-153) and that the helix is
stabilized on PrP-sen by intra-helix salt bridges between two aspartic
acid-arginine ion pairs at positions 144 and 148 and at 147 and 151, respectively. Mutants of the hamster prion protein were constructed by
replacing the aspartic acids with either asparagines or alanines to
destabilize the proposed helix 1 salt bridges. Thermal and chemical
denaturation experiments using circular dichroism spectroscopy
indicated the overall structures of the mutants are not substantially
destabilized but appear to unfold differently. Cell-free conversion
reactions performed using ionic denaturants, detergents, and salts
(conditions unfavorable to salt bridge formation) showed no significant
differences between conversion efficiencies of mutant and wild type
proteins. Using conditions more favorable to salt bridge formation, the mutant proteins converted with up to 4-fold higher efficiency than the
wild type protein. Thus, although spectroscopic data indicate the salt
bridges do not substantially stabilize PrP-sen, the cell-free
conversion data suggest that Asp-144 and Asp-147 and their
respective salt bridges stabilize PrP-sen from converting to
PrP-res.
Transmissible spongiform encephalopathies
(TSEs)1 or prion diseases are
neurological disorders characterized by the accumulation of abnormal
protease-resistant forms of prion protein (PrP), e.g. scrapie-associated PrP (PrPSc) or protease-resistant PrP
(PrP-res), in the central nervous system of diseased animals. TSE
diseases include Creutzfeldt-Jakob disease;
Gerstmann-Sträussler-Scheinker syndrome and kuru in humans;
scrapie in sheep, mice, and hamsters; bovine spongiform encephalopathy
in cattle; and chronic wasting disease in deer and elk. PrP-res
accumulates as a result of the conversion of the normal,
protease-sensitive form of PrP (PrPC or PrP-sen), which
occurs as a post-translational process without any apparent requirement
for covalent modifications (1, 2). Although not yet fully understood,
the accumulation of PrP-res appears to lead to cerebral amyloidosis,
dementia, motor dysfunction, and eventually death. PrP-res forms
insoluble aggregates and is partially resistant to proteinase K (PK),
which removes ~67 amino acid residues (6-7 kDa) from the N terminus
(1, 3-5). These differences in biophysical properties most likely
reflect different conformations of the two protein isoforms. Optical
spectroscopic data indicate that PrP-sen isolated from normal brain is
highly The conversion of PrP-sen into PrP-res is likely to be the key event in
the pathogenesis of TSE diseases. Although TSE diseases may be
sporadic, genetic, or infectious in origin, it is commonly argued that,
in each circumstance, the disease propagates itself through a
self-catalytic mechanism involving existing multimers of PrP-res. This
idea is supported by experiments that show that PrP-res can induce the
conversion of native PrP into PrP-res-like aggregates in
vitro (9, 10). However, the precise molecular mechanism of the
conversion between the normal and pathogenic forms of prion proteins
remains poorly understood.
Prior to conversion to PrP-res, the structure of PrP-sen (see Fig. 1)
consists of a core domain (residues 125-228), containing three
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-helical, whereas PrP-res contains a large proportion of
-sheet structure (6-8).
-helices (helix 1, residues 144-154; helix 2, 172-193; and helix
3, 200-227) and two short antiparallel
-strands (residues 129-131
and 161-163), and a largely unstructured N-terminal domain (residues
23-119) (11). Several considerations (12-15) suggest that helix 1 (residues 144-153) might be a site of interaction of PrP-sen and
PrP-res and that it plays a critical role in the initial stages of
PrP-sen unfolding and conversion. Morrissey and Shakhnovich (12) noted
that helix 1 is unusually hydrophilic and lacking in non-covalent
contacts with the remainder of the protein. They proposed that the
stabilization of helix 1 is self-sustaining through the presence of
intra-helix electrostatic interactions such as the two properly phased
Asp-Arg pairs within the helix, which form strong salt bridges and
stabilize the helical turns (Fig. 1)
(12). The directional ordering of the charges is expected to interact
with the intrinsic dipole moment of the helix providing further
stabilization. Molecular modeling studies suggested that the conserved
amino acid sequence of hamster PrP helix 1 might favorably adopt
conformations necessary to form intermolecular
-sheet aggregates and
that these aggregates might be stabilized by intermolecular
salt bridges between the helix 1 Asp and Arg residues of adjacent
molecules (12). Consistent with this idea, Vorberg et al.
reported that the deletion of helix 1 from PrP-sen completely inhibited
its conversion to PrP-res (31). Other studies have shown that a
de novo
to
conformational change and aggregation of
recombinant human PrP occurs in the presence of 1 M
guanidine hydrochloride (GdnHCl) and at pH levels below 5.0 (13-15). If the non-ionic denaturant urea was used, then the
conversion also required NaCl. It is possible that the protonation of
Asp-144 and Asp-147 at low pH interrupts the salt bridges in helix 1 and compromises its stability. The presence of the additional
denaturant and salts may also help disrupt the overall structure and
stability of the folded protein and screen electrostatic charges that
stabilize helix 1.
View larger version (17K):
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Fig. 1.
Salt bridges proposed to stabilize helix 1 of
PrP-sen. NMR structure of hamster PrP-sen (residues
125-228) reported by James et al. (left) (11).
Enlarged areas show helix 1 with stabilizing salt bridges
between aspartic acid 147 and arginine 151 (center) and side
chain atoms involved in the ionic interactions
(right).
To address the question of the role of helix 1 Asp residues and their
salt bridges to Arg residues in the stabilization of PrP-sen and its
conversion to PrP-res, we have prepared six mutants replacing Asp-144
and Asp-147 with either asparagines or alanines to eliminate the
putative salt bridges. The results indicate that PrP-sen is not
substantially stabilized overall by the Asp-Arg salt bridges and that
intermolecular salt bridge formation involving the same residues is not
essential for the formation of PrP-res. The unfolding pathway of
PrP-sen and its conversion to PrP-res, however, are influenced by
interactions involving the helix 1 aspartic acid residues. These
results provide insight into the basic mechanism of PrP-res formation.
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EXPERIMENTAL PROCEDURES |
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Prokaryotic Expression of Wild Type and Mutant Hamster PrP-(23-231)-- The plasmid encoding Syrian hamster PrP (HaPrP) (residues 23-231) was constructed in a pET24a(+) vector from Novagen inserted as an NdeI fragment using standard cloning techniques. Mutations at positions 144, 147, and both 144 and 147 were introduced using the QuikChange site-directed mutagenesis kit from Stratagene. Oligonucleotides used in the mutagenesis were obtained from Genosys Biotechnologies and include the following: (D144N) 5'-GCATTTTGGCAATAACTGGGAGGACCGCTACTACCG-3' and 5'-CGGTAGTAGCGGT CCTCCCAGTTATTGCCAAAATGC-3'; (D147N) 5'-GCAATGACTGGGAGAACCGCTACTACCGTGAAAAC-3' and 5'-GTTTTCACGGTAGTAGCGGTTCTCCCAGTCATTGC-3'; (D144N/D147N) 5'-GCATTTT GGCAATAACTGGGAGAACCGGTACTACCG-3' and 5'-CGGTAGTAGCGGTTCTCCCAGTTATTGCCAAAATGC-3'. Mutant DNA was transformed into XL1-Blue supercompetent cells from Stratagene. Mutant DNA was then purified from overnight cultures using a Wizard Miniprep plasmid purification kit. All constructs were sequenced by MWG Biotech. Plasmids were transformed into BL21(DE3) supercompetent cells from Stratagene for PrP protein expression. Recombinant proteins accumulated in cytoplasmic inclusion bodies.
Eukaryotic Expression of Wild Type and Mutant HaPrP--
D-to-N
mutant HaPrP (residues 23-231) in the pET24a(+) vector were subcloned
using Bsu36I and StuI restriction sites into full-length HaPrP without the glycosylphosphatidylinositol (GPI) anchor
in a pBluescript vector generously provided by Dr. Bruce Chesebro. The
D-to-A mutants were generated by site-directed mutagenesis of the HaPrP
pBluescript clone using a QuikChange site-directed mutagenesis kit from
Stratagene. Oligonucleotides were obtained from Genosys Biotechnologies
and include the following: (D144A) 5'-GCATTTTGGCAATGCCTGGGAGGACCGCTACTACCG-3' and
5'-CGGTAGTAGCGGTCCTCCCAGGCATTGCCAAAATGC-3'; (D147A)
5'-GCAATGACTGGGAGGCCCGC TACTACCGTGAAAAC-3' and
5'-GTTTTCACGGTAGTAGCGGGCCTCCCAGTCATTGC-3'; (D144A/D147A)
5'-GCATTTTGGCAATGCCTGGGAGGCCCGCTACTACCG-3' and
5'-CGGTAGTAGCGGGCCTCCCAGGCATTGCCAAAATGC-3'. The pBS HaPrP
construct lacks the GPI anchor, because the gene lacks the
C-terminal GPI attachment sequence (9). The pBS PrP clones were then
sequenced with an Applied Biosystems 3700 automated sequencer and
subcloned using standard techniques into the pSFF retroviral expression
vector using the EcoRI restriction site (16).
Mixtures of PA317 and 2 mouse fibroblast cells were transfected with
the pSFF plasmid, including the respective wild type or mutant PrP
insert and used for protein expression as described previously (9, 17,
18).
Prokaryotic Protein Expression and Purification--
Cells of
Escherichia coli BL21(DE3) with the desired PrP construct
were grown overnight in 5 ml of NZY liquid culture medium containing
0.05 mg/ml kanamycin. The overnight culture was added to 500 ml of NZY
medium with 0.05 mg/ml kanamycin and grown at 37 °C in a shaking
incubator to an optical density at 600 nm of 0.5. Protein expression
was then induced by adding isopropyl--D-thiogalactoside to 0.5 mM to the culture. The induced culture was grown for
2 h at 37 °C in a shaking incubator. Cells were then harvested
by centrifugation.
Cell pellets were suspended in lysis buffer (50 mM
Tris/HCl, pH 7.5, 2 mM EDTA, 0.1% Triton X-100) and
sonicated to release the inclusion bodies containing the protein of
interest. Inclusion bodies were collected by centrifugation and then
solubilized for 2 h in 8 M urea buffered at pH 8 with
0.1 M sodium phosphate and 0.01 M Tris. The
solubilized protein solution was then diluted 10-fold with 8.0 M urea in TCB buffer (20 mM Tris/HCl, pH 8.0, 150 mM NaCl, 2.5 mM CaCl2). The
solubilized protein solution was purified using Novagen His-Bind resin.
The resin was incubated with the solubilized protein in 8 M
urea for 2 h. The protein-bound resin was loaded onto the column
and washed with a 2 M step gradient from 8 to 0 M urea in TCB. The protein was eluted using 60 mM imidazole in TCB followed by 200 mM
imidazole in TCB. Fractions were analyzed using SDS-PAGE to confirm the
presence of PrP (molecular mass, 23-kDa). Fractions containing
only PrP were pooled and dialyzed against 0.01 M sodium
phosphate at pH 6.0 to remove the imidazole and then concentrated using
a centrifugal filter device (molecular mass cut-off, 10 kDa) to ~0.5
mg/ml final protein concentration (280 = 2.7 ml
mg
1 cm
1). No impurities were seen in the
final protein preparations using SDS-PAGE and Coomassie Blue staining.
Molecular masses for all purified protein constructs were confirmed
using MALDI mass spectrometry. The molecular masses for the D144N,
D147N, and D144N/D147N mutants were 0.95, 0.82, and 1.89 atomic mass
units lower, respectively, than the wild type protein, consistent with
the respective theoretical molecular masses.
Circular Dichroism Spectroscopy and Denaturation Curves-- Circular Dichroism (CD) spectroscopy was performed using an On-line Instrument Systems (Olis) CD module with a Cary-16 spectrophotometer conversion unit with a NESLAB RTE-111 temperature control module. For thermal denaturation experiments the ellipticity from 225 to 207.5 nm of PrP samples at 0.3 mg/ml in 0.01 M NaPO4 at pH 6.0 was measured at intervals between 25 and 72.5 °C. Full spectra at 25, 62.5, and 72.5 °C were scanned from 185 to 260 nm. Samples were scanned three times with a 0.5-nm scanning interval in a 1.0-mm cylindrical cell after equilibrating 15 min at the desired temperature. Scans were averaged using the Olis spectral processing software supplied with the instrument. The fraction of unfolded protein for each construct was determined using the molar ellipticity at 222 nm and was plotted versus the temperature in degrees Celsius. The ellipticity at 72.5 °C was used for the unfolded protein ellipticity. All of the mutant protein signals were normalized to the wild type PrP unfolded signal. Averages and standard deviations of transition midpoints and slopes were determined from three individual experiments for each protein.
For the GdnHCl denaturation experiments, samples from 0 to 5.0 M GdnHCl were prepared by diluting a stock PrP solution
(0.3 mg/ml) with varying amounts of a stock 8.0 M GdnHCl
solution and buffer giving a final protein concentration of 0.1 mg/ml.
All solutions were buffered at pH 6.0 in 0.01 M
NaPO4. Samples were allowed to equilibrate for at least
1 h prior to analysis. Samples were scanned three times in a
1.0-mm cylindrical cell from 225 to 207.5 nm with a 0.5-nm scanning
interval at 25 °C. All spectral data were corrected for absorbance
by the buffer and denaturant. The fraction of unfolded protein was
determined for each construct using the ellipticity at 222 nm. Averages
and ranges for the thermodynamic parameters were determined from two
individual experiments for each protein. Thermal and GdnHCl
denaturation curves were fit using the non-linear regression analysis
tool on GraphPad Prism. Thermal unfolding transition midpoints and
GdnHCl denaturation thermodynamic parameters were determined assuming a
two-state unfolding process as described by Pace and Scholtz (19).
Plots of G0 versus GdnHCl
concentration were fit by linear regression according to the equation,
G0 =
G(H2O)
m[GdnHCl], where
G0 is the
standard Gibbs free energy,
G(H2O) is the
Gibbs free energy of unfolding in the absence of denaturant,
m is the cooperativity of unfolding, and [GdnHCl] is the
concentration of GdnHCl in molarity.
Metabolic [35S]Methionine Labeling of Wild Type and
Mutant PrP-sen--
Metabolic labeling of PA317 and 2 mouse
fibroblast cells with the wild type or mutant PrP pSFF clones was
performed as described previously (20-22). Lysis and extraction of PrP
from cells was then performed as described previously (23, 24). The
protein was immunoprecipitated using the mouse 3F4 monoclonal antibody as described previously (24).
Cell-free Conversion Reaction--
PrP-res from 263K
scrapie-infected hamsters was prepared using a revision of the Bolton
et al. method (25) and was stored in 0.5% sulfobetaine
(Calbiochem, La Jolla, CA, Zwittergent 3-14) in phosphate-buffered
saline as described previously (9, 20). Crude normal and 263K-infected
hamster brain microsomes were prepared as described previously (26).
Two different cell-free reaction conditions were used: 1) reactions
using purified PrP-res pretreated with 2 M GdnHCl for
1 h at 37 °C and 2) reactions using PrP-res in brain
microsomes. Cell-free conversion reactions in the presence of GdnHCl
using purified PrP-res and reactions without detergents and denaturants
using microsomal PrP-res were performed as described previously (9, 21,
24, 26). Each reaction contained 100 ng of 263K PrP-res mixed with
10,000 to 20,000 cpm of [35S]PrP-sen. For reactions using
GdnHCl pretreated PrP-res final reaction mixtures also contained 364 mM GdnHCl, 34 mM Sarkosyl, 1.25 mM
sulfabetaine, 5 nM hexadecylpyridinium chloride, and
50 mM sodium citrate at pH 6.0. For conversions using
microsomal PrP-res, final reaction mixtures also contained 100 mM NaCl, 5 mM MgCl2, and 50 mM sodium citrate at pH 6.0. Reactions were incubated for
40-48 h at 37 °C. After incubation, 90% of the reaction mixture was treated with 20 µg/ml PK at 37 °C for 1 h. The remaining
10% was analyzed without PK treatment. The PK digestion was stopped by
adding Pefabloc (Roche Molecular Biochemicals) to 2 mM, 20 µg of thyroglobulin as a carrier protein, and four volumes of methanol. Precipitated proteins were collected by centrifugation and
separated by SDS-PAGE using Novex NuPAGE pre-cast polyacrylamide gels.
Radioactive proteins were visualized and quantified using a
PhosphorImager (Amersham Biosciences).
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RESULTS |
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Disulfide Bond Formation and Protein Purity--
To investigate
the roles of the helix 1 Asp residues and their proposed salt bridges
on the stability of the secondary structure of PrP, three mutant
proteins (D144N, D147N, and D144N/D147N) of full-length hamster PrP
(residues 23-231) were expressed as cytoplasmic inclusion bodies in
Escherichia coli, purified, and refolded for analysis by
circular dichroism spectroscopy. The purified proteins were analyzed by
SDS-PAGE to assess protein purity and the presence of the disulfide
bond between Cys179 and Cys 214 (Fig. 2).
The slower migration of the proteins on SDS-PAGE upon treatment with
-mercaptoethanol (lanes 1-4) than untreated samples
(lanes 5-8) indicated that the disulfide bond was present in the untreated preparations of each construct (27). Each protein ran
as a single band with an estimated purity of at least 90%.
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Thermal Denaturation Monitored by CD Spectroscopy--
The
secondary structures and stabilities of the PrP mutants relative to
wild type PrP were assessed by CD spectroscopy. Under non-denaturing
conditions at 25 °C and pH 6.0 (the optimal pH for most
PrP-res-induced cell-free conversion reactions), spectra of the wild
type and mutant PrP constructs (Fig.
3a) were identical, each
showing negative lobes at 208 and 222 nm indicative of highly -helical secondary structures as has been seen previously for PrP-sen derived from brain (6), mammalian tissue culture cells (28),
and recombinant PrP synthesized and refolded from inclusion bodies
(29). These data suggest the Asp residues and their respective salt
bridges do not substantially stabilize helix 1. Thermal denaturation was performed to determine if the mutants were destabilized with respect to secondary structure and unfolding transitions. The thermal
denaturation curves were sigmoidal for all constructs, suggesting that
they undergo a two-state unfolding transition (Fig. 3b).
However, none of the thermal unfolding reactions for the PrP constructs
were reversible; thus, thermodynamic values could not be obtained.
Transition midpoints and slopes of the unfolding transition regions
were used for comparison instead. The transition midpoints (Table
I) for the mutants were within 2.2 °C
of the wild type protein transition midpoint suggesting that the PrP
secondary structure is not substantially destabilized by the mutations
and the presumed elimination of the helix 1 salt bridges. However, the
slopes of the transition regions of the mutant unfolding curves (Table
I) were lower than the slope of the wild type protein unfolding curve.
These data indicate that the mutants unfold differently and less
cooperatively than the wild type protein.
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To look more closely at the thermal denaturation differences between
the wild type and mutant PrP molecules, CD spectra at 62.5 °C,
72.5 °C, and, 25 °C after cooling from 72.5 °C, were overlaid
(Fig. 3, c-e). The wild type protein changed from a highly -helical conformation at 25 °C (Fig. 3a) to a highly
-sheet conformation at 62.5 and 72.5 °C as indicated by the
negative peak centered at 215 nm (Fig. 3, c and
d). At 62.5 °C all three mutants appeared to still have
some
-helical content judging by the slight negative dips in
ellipticity at 208 and 222 nm (Fig. 3c). At 72.5 °C the
mutant spectra all had negative lobes around 204 nm different from the
wild type spectrum, most noticeable in the D144N and D144N/D147N mutant
spectra (Fig. 3d). This difference indicated that the mutant
proteins have different states of denaturation at 72.5 °C than the
wild type. Negative ellipticities below 200 nm are associated with
disordered structure; thus, the mutant PrP conformations appeared to
contain more disordered structure than the more
-sheet-containing
wild type protein. In addition to the conformational differences at
72.5 °C, there were differences in the reversibilities of the
unfolding induced by this temperature. Thermal denaturation of the wild
type protein was highly irreversible, which has been shown previously
(30), as can be seen by the fact that the spectra at 72.5 and 25 °C
after cooling from 72.5 °C are shaped similarly with main negative
lobes for both spectra centered close to 215 nm (Fig. 3, d
and e). Spectra of the mutant PrP molecules after cooling
from 72.5 °C to 25 °C showed a partial change back to more
-helical spectra with the D144N and D144N/D147N spectra, in
particular, showing negative lobes at 208 and 222 nm (Fig.
3e), which are similar to, but not identical to the spectra at 25 °C prior to thermal denaturation (Fig. 3a). These
data suggest the mutant proteins unfold to a different denatured state
structure at 72.5 °C than the wild type protein.
GdnHCl Denaturation Monitored by CD Spectroscopy--
To further
investigate the unfolding of the PrP helix 1 mutants, GdnHCl unfolded
samples of wild type D144N, D147N, and D144N/D147N PrP at pH 6.0 were
monitored using CD spectroscopy at 222 nm. As shown in Fig.
4, the sigmoid shapes of the unfolding
curves indicated that all the constructs underwent a cooperative
two-state transition. All unfolding reactions were found to be
reversible (data not shown). Thermodynamic parameters, determined from
the unfolding data assuming a two-state transition, are shown in Table I. The wild type protein had a free energy of conformation
((G(H2O)) similar to that for recombinant
full-length ovine PrP (27) but lower than that reported for full-length
murine PrP (14). The mutant proteins have lower
G(H2O) values and transition midpoints ([GdnHCl]1/2) than wild type PrP. The cooperativities for the
unfolding (m) indicated that the wild type
(m = 7.86 kJ/mol/M), D144N
(m = 7.43 kJ/mol/M), and D147N (m = 7.85 kJ/mol/M) PrP unfolded more
cooperatively than did the D144N/D147N (m = 6.04 kJ/mol/M) mutant. The
G(H2O) and
[GdnHCl]1/2 data support the conclusion from the thermal
denaturation experiments that the mutants unfold somewhat differently
than the wild type protein. However, although the mutants tended to be
slightly more stable against thermal denaturation, they were slightly
less stable than the wild type protein to GdnHCl-induced denaturation.
Nonetheless, neither the thermal nor the chemical denaturation data
suggest that the overall stability of PrP is substantially dependent
upon the proposed helix 1 Asp residues and their salt bridges.
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Cell-free Conversion--
It remained to be determined whether the
helix 1 aspartic acid residues are involved in the interactions
necessary for the conversion of PrP-sen to PrP-res. Specifically, if
the critical interaction required for the conversion involved
intermolecular ionic bonds between aspartic acid (Asp-144 and Asp-147)
and arginine (Arg-148 and Arg-151) residues as proposed (12), one would
expect conversion of the mutants lacking the Asp residues to be
inhibited. To address this question the D144N, D147N, and D144N/D147N
PrP mutants were expressed in mammalian cells, radiolabeled using [35S]methionine, and analyzed using the cell-free
conversion assay (9). The assay is performed by "seeding" the
conversion of 35S-labeled PrP-sen using PrP-res from the
brains of hamsters infected with 263K scrapie. After an incubation
period, newly converted (radiolabeled), partially protease-resistant
PrP is detected following digestion with proteinase K by SDS-PAGE and
phosphor autoradiography (9, 10). We limited the conversion reactions
to PrP-sen that was metabolically radiolabeled in mammalian cells
because of difficulties in generating [35S]methionine
labeled PrP-sen with sufficient specific activity when expressed in
E. coli. Fig. 5 shows the
results of conversion reactions performed in the presence of detergents
and GdnHCl using purified 263K hamster PrP-res to drive the reaction.
Fig. 5a shows the undigested radiolabeled PrP
post-incubation without (lanes 1-4) and with (lanes
5-8) PrP-res. Lanes 9-16 show the remaining portion
of the same samples after a 1-h digestion with PK. The lanes incubated
without PrP-res had no protease-resistant bands. However, samples
incubated in the presence of PrP-res showed distinct protease-resistant
35S-PrP bands that, like PK digestion products of
brain-derived PrP-res itself, were 6-7 kDa lower in molecular mass
than full-length precursor 35S-PrP-sen molecules.
Quantification of the proportion of the input 35S-PrP-sen
converted to PK-resistant bands of 19-26 kDa (i.e.
conversion efficiency) is shown in Fig. 5b. In this case all
three mutants converted (lanes 14-16) with statistically
indistinguishable efficiencies from the wild type PrP control
(lane 13). These data are not consistent with the previously
observed conversion inhibition of PrP helix 1 deletion mutants (31),
suggesting that aspartic acid residues 144 and 147 are not essential
for the conversion of PrP-sen to PrP-res in the presence of detergents
and GdnHCl.
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Because the above conversion conditions (especially the presence of
GdnHCl) may have blocked the ionic interactions necessary for the
presence of the helix 1 salt bridges, conversion reactions were
performed without detergents or denaturants as described recently (10).
This method uses a crude scrapie brain microsome fraction containing
PrP-res to drive conversion. These conversion reactions more closely
mimic conditions found on cell membranes where PrP-sen is located and
most likely converted in scrapie-infected cells (23, 32). Under these
conditions, the PrP mutants converted with two-three times the
conversion efficiency as the wild type PrP (Fig.
6, a and b). Thus,
the replacement of the aspartic acid residues with asparagines at
positions 144 and 147 enhanced, rather than prevented, conversion to
PrP-res.
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The influence of the helix 1 salt bridges on PrP conversion was also
tested using 35S-labeled PrP-sen mutants where the aspartic
acids at positions 144 and 147 were individually or both replaced with
alanines. These replacements should have prevented ion-dipole
interactions in addition to ion-ion interactions between the side
chains of the residues involved in the helix 1 salt bridges in wild
type PrP. If the increases in conversion efficiencies of the D-to-N PrP
mutants were solely due to the loss of a salt bridge or any other
interaction that stabilizes the native -helical structure from
converting to PrP-res, then one would expect that the D-to-A mutant
proteins would show the same conversion behavior as the respective
D-to-N mutants. Using radiolabeled D144A, D147A, and D144A/D147A
PrP-sen mutants in the detergent and GdnHCl-free conversion reactions,
we produced the expected conversion products (Fig. 7a, compare lanes
9-16 to lanes 1-8). The conversion efficiencies of
the D-to-A mutant PrP constructs were roughly three to four times as
efficient as the conversion efficiency of the wild type PrP (Fig.
7b), confirming that mutations expected to disrupt the helix
1 salt bridges promote PrP-res-induced conversion. The fact that D-to-A
mutants were even more efficiently converted than the D-to-N mutants
suggests that, in the latter, some stabilizing interactions between the
Asn-144, Asn-147, and the arginines in helix 1 (presumably ion-dipole
interactions) still occur.
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DISCUSSION |
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This investigation sought to determine the role of the putative Asp-Arg helix 1 salt bridges in the stability of PrP-sen and whether the same residues proposed to be involved in the salt bridges play a role in the conversion of PrP-sen to PrP-res. It appears from our data that helix 1 does not behave as has been previously proposed (12). That is, our data suggest that the helix 1 salt bridges do not substantially stabilize PrP under native conditions, and, intermolecular salt bridge formation between the same salt bridge residues is not critical for the conversion of PrP-sen to PrP-res. Instead it appears that the aspartic acid residues affect how the protein unfolds and provide stability against the conversion of PrP-sen to PrP-res.
The contribution of surface salt bridges, such as those proposed to exist on helix 1, has generally been considered to contribute little to the overall stability of protein structures. It has been reported that salt bridges buried in the protein interior contribute as much as 20.9 kJ/mol, whereas surface salt bridges contribute only 2.1-5.0 kJ/mol to protein-free energies of conformation (33, 34). Our data are consistent with this consensus, because the Gibbs free energies of conformation of the D144N, D147N, and D144N/D147N mutants were estimated by GdnHCl unfolding to be only 2.3, 0.9, and 5.1 kJ/mol lower, respectively, than wild type hamster PrP-sen. These data, combined with the fact the CD spectra of the mutants are identical to the wild type protein at 25 °C, suggest that there is no unusual stability gain due to ionic interactions involving the aspartic acid and arginine residues of helix 1 of hamster PrP-sen.
Although the effects of the helix 1 mutations on the overall free energy of PrP are modest, the localized effects on helix 1 stability might be more dramatic. At pH levels below 4.0, three-state unfolding transitions have been reported for recombinant human PrP in the presence of GdnHCl and urea with salt (30, 35). It is tempting to think that the first transition might involve the unfolding of a localized domain containing helix 1, because this helix has been modeled to lose helical structure at low pH (36) and has been shown to more easily undergo hydrogen-deuterium exchange than the central portions of helices 2 and 3 (37), suggesting a looser, less compact structure that might unfold more readily. Such a transition at low pH might be dependent on the protonation and ionic screening of the aspartic acids disrupting the salt bridges leading to the unfolding of helix 1 prior to the unfolding of the remaining helices. If such a three-state transition were occurring under the conditions of our experiments (i.e. pH 6), one might expect to have detected it in our unfolding curves monitored at 222 nm, because helix 1 makes up 18% of the total helix content of PrP-sen. However, our unfolding curves of both the wild type and helix 1 mutants suggested only two-state processes without any additional inflection points. Nonetheless, the helix 1 mutations reduced the slopes of the transition regions for the thermal denaturation curves, indicating the mutants unfold less cooperatively than the wild type protein. This effect was especially apparent for the D144N/D147N double mutant. Presumably, therefore, replacement of the helix 1 aspartates destabilizes helix 1 to some extent, but our data provide no evidence that this allows helix 1 to unfold independently of the other helices under the conditions suitable for the PrP-res-induced conversion reaction. One possibility is that helix 1 is already partially unfolded under these conditions, reducing our chances of detecting any independent unfolding in the denaturation curves. In any case, the low structural resolution of CD spectroscopy prevents us from fully discerning the conformational effects of the aspartate replacement and elimination of putative helix 1 salt bridges.
The differences between unfolding cooperativities of the mutant and wild type PrP molecules raise the possibility that the helix 1 aspartic acids are involved with stabilizing interactions other than the intra-molecular salt bridges. Eleven different salt bridges involving seven different anionic residues and eight different cationic residues were predicted in molecular dynamic simulations of human and Syrian hamster PrP-sen as reported by Zuegg and Gready (38). Five of the eleven proposed salt bridges involve either Asp-144 or Asp-147, including, in addition to the intra-helix salt bridges investigated here, Asp-144 with His-140, Asp-144 with Arg-208, and Asp-147 with His-140. The differences in unfolding cooperativities could result from the elimination of any of the five proposed salt bridges.
Protein unfolding cooperativity differences can also result from differences in the stabilities of the denatured state conformations (33, 39). The PrP thermal denaturation data show that the denatured states of the mutants at 72.5 °C differ from the wild type protein suggesting the mutants unfold via somewhat different conformations. Such alterations in the unfolding pathway, rather than differences in overall thermodynamic stability, could account for the increased conversion efficiencies seen in our cell-free conversion experiments. Consistent with this possibility are observations that decreased thermodynamic stability of PrPC is not a common characteristic of mutants associated with familial Creutzfeldt-Jakob disease, Gerstmann-Sträussler-Scheinker syndrome, and fatal familial insomnia (40, 41). Reasons proposed for how mutations associated with inherited forms of spongiform encephalopathies lead to disease include: (a) an increase in the stability of PrPSc; (b) an increased fraction of alternative conformations in equilibrium with PrPC, which may act as precursors to PrPSc; and (c) differences in PrPC folding kinetics with accumulation of PrPSc-like kinetic folding intermediates (41). Any of these reasons could apply as reasons for the increase in PrP-res formation seen in the cell-free conversion reactions using microsomal PrP-res and the PrP-sen helix 1 mutants studied herein.
Thus, the helix 1 mutants may populate more convertible conformations
along the PrP-sen to PrP-res conversion pathway to a greater extent
than wild type PrP. Alternatively, because the helix 1 mutants do not
unfold to the same -sheet-containing denatured structures at 62.5 and 72.5 °C as the wild type protein, it is possible that the
mutants populate an off-pathway
-sheet conformation to a lesser
extent than the wild type protein. The formation of an off-pathway
-oligomer has been previously reported to occur during amyloid
fibril formation of recombinant PrP 27-30 (42).
The more disordered conformations seen with the helix 1 mutants during
thermal denaturation might be more convertible to PrP-res than the more
-sheet conformation of wild type PrP. However, Rezaei et
al. (43) recently reported that the thermally denatured conformations of ovine PrP variants associated with low susceptibility to scrapie infection contained more disordered structure, whereas the
highly susceptible variants contained more
-sheet. The ovine PrP
variant data is not consistent with our data, because our mutants show
more disordered structure but convert with higher efficiencies relative
to the wild type protein. Thus, it appears that unfolding to a more
disordered conformation is not necessarily associated with increased
susceptibility to PrP-res formation. Rather, our data suggest that our
D-to-N mutants populate intermediate conformations that contain
unspecified secondary structures with less
-sheet that are more
susceptible to PrP-res formation than wild type PrP.
Considering all the data, three mechanistic pathways leading to PrP-res
formation can be proposed assuming the conversion is based on the
autocatalytic seeded polymerization model (40, 44, 45) (Fig.
8). Assuming PrP-sen is in equilibrium
with convertible, intermediate, and non-convertible conformations, the
pathways vary depending on which conformation binds to PrP-res. In
pathway a, the intermediate PrP-sen conformation binds to PrP-res. After binding, the PrP-sen molecule can undergo a conformational transition to either the convertible conformation or to the off-pathway non-convertible conformation. PrP-res is only formed when the bound
PrP-sen molecule attains the convertible conformation. In pathway b,
convertible PrP-sen binds to PrP-res and then can convert directly to
PrP-res. In pathway c, a non-convertible PrP-sen conformer binds to
PrP-res and must transform to intermediate and/or convertible PrP-sen
prior to converting to PrP-res. Evidence for the existence of bound,
but non-convertible states has come from previous studies of
interspecies conversion reactions (46). In the case of the helix 1 mutants, PrP-sen is proposed to be less likely to adopt the
non-convertible conformation resulting in more conversion of the mutant
PrP-sen molecules to PrP-res than the wild type PrP-sen.
|
In summary, these results suggest that PrP-sen is not substantially
stabilized by the aspartic acid-arginine salt bridges. However, the
helix 1 mutants unfold less cooperatively and do not attain the same
-sheet denatured state as the wild type protein upon thermal
denaturation. Cell-free conversion data suggest the salt bridge
residues are not critical for the formation of PrP-res, because
protease-resistant material was generated in all conversion reactions
performed with and without the presence of detergents and denaturants.
The conversion efficiencies of the mutants in the absence of detergents
and denaturants, conditions favorable to salt bridge formation, are
higher than the wild type protein, indicating that the elimination of
the anionic charge of the aspartic acids increases the convertibility
of PrP-sen. The increase in conversion could possibly be due to the
elimination of an off-pathway conformation along the conformational
pathway leading to conversion to PrP-res.
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ACKNOWLEDGEMENTS |
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We thank S. Fyffe for generating the recombinant mutant PrP E. coli clones and G. Baron, G. Raymond, S. Priola, L. Raymond, G. Sylva, I. Vorberg, J. Wolfinbarger, and K. Wehrly for additional technical assistance. We also thank A. Robertson, R. Swanstrom, and M. McGuirl for helpful discussions, G. Hettrick for graphics assistance, A. Hughson for the MALDI mass spectrometry analysis and L. Evans, R. Kodali, and S. Sampath for critical reading of the manuscript.
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FOOTNOTES |
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* The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
¶ Current address: Dept. of Biological Chemistry, Wyeth Research, 85 Bolton St., Cambridge, MA 02140.
To whom correspondence should be addressed. Tel.:
406-363-9264; Fax: 406-363-9286; E-mail: bcaughey@nih.gov.
Published, JBC Papers in Press, January 27, 2003, DOI 10.1074/jbc.M211599200
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ABBREVIATIONS |
---|
The abbreviations used are:
TSE, transmissible
spongiform encephalopathy;
PrP, prion protein;
PrPSc, scrapie-associated prion protein;
PrP-res, protease-resistant prion
protein;
PrPC, cellular prion protein;
PrP-sen, protease-sensitive prion protein;
PK, proteinase K;
GdnHCl, guanidine
hydrochloride;
HaPrP, Syrian hamster prion protein;
GPI, glycosylphosphatidylinositol;
CD, circular dichroism;
G(H2O), free energy of conformation;
[GdnHCl]1/2, guanidine unfolding transition midpoint;
m, cooperativity of unfolding in guanidine hydrochloride;
MALDI, matrix-assisted laser desorption ionization.
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REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Hope, J., Morton, L. J., Farquhar, C. F., Multhaup, G., Beyreuther, K., and Kimberlin, R. H. (1986) EMBO J. 5, 2591-2597[Abstract] |
2. | Stahl, N., Baldwin, M. A., Teplow, D. B., Hood, L., Gibson, B. W., Burlingame, A. L., and Prusiner, S. B. (1993) Biochemistry 32, 1991-2002[Medline] [Order article via Infotrieve] |
3. | Oesch, B., Westaway, D., Walchli, M., McKinley, M. P., Kent, S. B., Aebersold, R., Barry, R. A., Tempst, P., Teplow, D. B., and Hood, L. E. (1985) Cell 40, 735-746[Medline] [Order article via Infotrieve] |
4. | Meyer, R. K., McKinley, M. P., Bowman, K. A., Braunfeld, M. B., Barry, R. A., and Prusiner, S. B. (1986) Proc. Natl. Acad. Sci. U. S. A. 83, 2310-2314[Abstract] |
5. | Rubenstein, R., Kascsak, R. J., Merz, P. A., Papini, M. C., Carp, R. I., Robakis, N. K., and Wisniewski, H. M. (1986) J. Gen. Virol. 67, 671-681[Abstract] |
6. | Pan, K. M., Baldwin, M., Nguyen, J., Gasset, M., Serban, A., Groth, D., Mehlhorn, I., Huang, Z., Fletterick, R. J., and Cohen, F. E. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 10962-10966[Abstract] |
7. | Caughey, B. W., Dong, A., Bhat, K. S., Ernst, D., Hayes, S. F., and Caughey, W. S. (1991) Biochemistry 30, 7672-7680[Medline] [Order article via Infotrieve] |
8. |
Safar, J.,
Roller, P. P.,
Gajdusek, D. C.,
and Gibbs, C. J., Jr.
(1993)
Protein Sci.
2,
2206-2216 |
9. | Kocisko, D. A., Come, J. H., Priola, S. A., Chesebro, B., Raymond, G. J., Lansbury, P. T., and Caughey, B. (1994) Nature 370, 471-474[CrossRef][Medline] [Order article via Infotrieve] |
10. | Caughey, B., Raymond, G. J., Callahan, M. A., Wong, C., Baron, G. S., and Xiong, L. W. (2001) Adv. Protein Chem. 57, 139-169[CrossRef][Medline] [Order article via Infotrieve] |
11. |
James, T. L.,
Liu, H.,
Ulyanov, N. B.,
Farr-Jones, S.,
Zhang, H.,
Donne, D. G.,
Kaneko, K.,
Groth, D.,
Mehlhorn, I.,
Prusiner, S. B.,
and Cohen, F. E.
(1997)
Proc. Natl. Acad. Sci. U. S. A.
94,
10086-10091 |
12. |
Morrissey, M. P.,
and Shakhnovich, E. I.
(1999)
Proc. Natl. Acad. Sci. U. S. A.
96,
11293-11298 |
13. | Swietnicki, W., Morillas, M., Chen, S. G., Gambetti, P., and Surewicz, W. K. (2000) Biochemistry 39, 424-431[CrossRef][Medline] [Order article via Infotrieve] |
14. |
Hornemann, S.,
and Glockshuber, R.
(1998)
Proc. Natl. Acad. Sci. U. S. A.
95,
6010-6014 |
15. | Kaneko, K., Peretz, D., Pan, K. M., Blochberger, T. C., Wille, H., Gabizon, R., Griffith, O. H., Cohen, F. E., Baldwin, M. A., and Prusiner, S. B. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 11160-11164[Abstract] |
16. | Chesebro, B., Wehrly, K., Caughey, B., Nishio, J., Ernst, D., and Race, R. (1993) Dev. Biol. Stand. 80, 131-140[Medline] [Order article via Infotrieve] |
17. | Robertson, M. N., Spangrude, G. J., Hasenkrug, K., Perry, L., Nishio, J., Wehrly, K., and Chesebro, B. (1992) J. Virol. 66, 3271-3277[Abstract] |
18. | Priola, S. A., Caughey, B., Race, R. E., and Chesebro, B. (1994) J. Virol. 68, 4873-4878[Abstract] |
19. | Pace, C. N., and Scholtz, J. M. (1997) in Protein Structure: A Practical Approach (Creighton, T. E., ed), 2nd Ed. , pp. 299-321, Oxford University Press, New York |
20. |
Kocisko, D. A.,
Priola, S. A.,
Raymond, G. J.,
Chesebro, B.,
Lansbury, P. T., Jr.,
and Caughey, B.
(1995)
Proc. Natl. Acad. Sci. U. S. A.
92,
3923-3927 |
21. |
Horiuchi, M.,
and Caughey, B.
(1999)
EMBO J.
18,
3193-3203 |
22. | Caughey, B., Horiuchi, M., Demaimay, R., and Raymond, G. J. (1999) Methods Enzymol. 309, 122-133[Medline] [Order article via Infotrieve] |
23. |
Caughey, B.,
and Raymond, G. J.
(1991)
J. Biol. Chem.
266,
18217-18223 |
24. | Caughey, B., Kocisko, D. A., Raymond, G. J., and Lansbury, P. T., Jr. (1995) Chem. Biol. 2, 807-817[Medline] [Order article via Infotrieve] |
25. | Bolton, D. C., Bendheim, P. E., Marmorstein, A. D., and Potempska, A. (1987) Arch. Biochem. Biophys. 258, 579-590[Medline] [Order article via Infotrieve] |
26. |
Baron, G. S.,
Wehrly, K.,
Dorward, D. W.,
Chesebro, B.,
and Caughey, B.
(2002)
EMBO J.
21,
1031-1040 |
27. |
Rezaei, H.,
Marc, D.,
Choiset, Y.,
Takahashi, M.,
Hui Bon, H. G.,
Haertle, T.,
Grosclaude, J.,
and Debey, P.
(2000)
Eur. J. Biochem.
267,
2833-2839 |
28. | Xiong, L. W., Raymond, L. D., Hayes, S. F., Raymond, G. J., and Caughey, B. (2001) J. Neurochem. 79, 669-678[CrossRef][Medline] [Order article via Infotrieve] |
29. | Hornemann, S., Korth, C., Oesch, B., Riek, R., Wider, G., Wuthrich, K., and Glockshuber, R. (1997) FEBS Lett. 413, 277-281[CrossRef][Medline] [Order article via Infotrieve] |
30. | Jackson, G. S., Hill, A. F., Joseph, C., Hosszu, L., Power, A., Waltho, J. P., Clarke, A. R., and Collinge, J. (1999) Biochim. Biophys. Acta 1431, 1-13[Medline] [Order article via Infotrieve] |
31. |
Vorberg, I.,
Chan, K.,
and Priola, S. A.
(2001)
J. Virol.
75,
10024-10032 |
32. | Caughey, B., Raymond, G. J., Ernst, D., and Race, R. E. (1991) J. Virol. 65, 6597-6603[Medline] [Order article via Infotrieve] |
33. | Matthews, B. W. (1993) Annu. Rev. Biochem. 62, 139-160[CrossRef][Medline] [Order article via Infotrieve] |
34. | Nakamura, H. (1996) Q. Rev. Biophys. 29, 1-90[Medline] [Order article via Infotrieve] |
35. |
Swietnicki, W.,
Petersen, R.,
Gambetti, P.,
and Surewicz, W. K.
(1997)
J. Biol. Chem.
272,
27517-27520 |
36. | Alonso, D. O., and Daggett, V. (2001) Adv. Protein Chem. 57, 107-137[Medline] [Order article via Infotrieve] |
37. | Liu, H., Farr-Jones, S., Ulyanov, N. B., Llinas, M., Marqusee, S., Groth, D., Cohen, F. E., Prusiner, S. B., and James, T. L. (1999) Biochemistry 38, 5362-5377[CrossRef][Medline] [Order article via Infotrieve] |
38. | Zuegg, J., and Gready, J. E. (1999) Biochemistry 38, 13862-13876[CrossRef][Medline] [Order article via Infotrieve] |
39. |
Shortle, D.
(1996)
FASEB J.
10,
27-34 |
40. | Griffith, J. S. (1967) Nature 215, 1043-1044[Medline] [Order article via Infotrieve] |
41. | Liemann, S., and Glockshuber, R. (1999) Biochemistry 38, 3258-3267[CrossRef][Medline] [Order article via Infotrieve] |
42. |
Baskakov, I. V.,
Legname, G.,
Baldwin, M. A.,
Prusiner, S. B.,
and Cohen, F. E.
(2002)
J. Biol. Chem.
277,
21140-21148 |
43. | Rezaei, H., Choiset, Y., Eghiaian, F., Treguer, E., Mentre, P., Debey, P., Grosclaude, J., and Haertle, T. (2002) J. Mol. Biol. 322, 799-814[CrossRef][Medline] [Order article via Infotrieve] |
44. | Come, J. H., Fraser, P. E., and Lansbury, P. T., Jr. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 5959-5963[Abstract] |
45. | Caughey, B. (2001) Trends Biochem. Sci. 26, 235-242[CrossRef][Medline] [Order article via Infotrieve] |
46. |
Horiuchi, M.,
Priola, S. A.,
Chabry, J.,
and Caughey, B.
(2000)
Proc. Natl. Acad. Sci. U. S. A.
97,
5836-5841 |