Download Heat stability of prion rods and recombinant prion protein in water

Journal
of General Virology (2001), 82, 465–473. Printed in Great Britain
...................................................................................................................................................................................................................................................................................
Heat stability of prion rods and recombinant prion protein in
water, lipid and lipid–water mixtures
Thomas Raul Appel,1† Michael Wolff,1 Friedrich von Rheinbaben,2 Michael Heinzel2
and Detlev Riesner1
1
2
Heinrich-Heine-Universita$ t Du$ sseldorf, Institut fu$ r Physikalische Biologie, Geba$ ude 26.12, D-40225 Du$ sseldorf, Germany
Henkel KGaA, Du$ sseldorf, Germany
Prion rods, i.e. insoluble infectious aggregates of the N-terminally truncated form of the prion
protein, PrP 27–30, and the corresponding recombinant protein, rPrP(90–231), were autoclaved
in water, bovine lipid or lipid–water mixtures for 20 min at temperatures from 100 to 170 SC. A
protocol was developed for the quantitative precipitation of small amounts of protein from large
excesses of lipid. PrP remaining undegraded after autoclaving was quantified by Western blot and
degradation factors were calculated. The Arrhenius plot of the rate of degradation vs temperature
yielded linear relationships for prion rods in water or lipid–water as well as for rPrP(90–231) in
lipid–water. The presence of lipids increased the heat stability of prion rods, especially at lower
temperatures. Prion rods had a much higher thermal stability compared to rPrP. Autoclaving of
prion rods in pure lipid gave different results – not simple degradation but bands indicative of
covalently linked dimers, tetramers and higher aggregates. The heat stability of prion rods in pure
lipid exceeded that in lipid–water mixtures. Degradation factors larger than 104 were reached at
170 SC in the presence of lipids and at 150 SC in the absence of lipids. The linear correlation of the
data allows cautious extrapolation to conditions not tested, i.e. temperatures higher than 170 SC.
A factual basis for assessing the biological safety of industrial processes utilizing potentially BSEor scrapie-contaminated animal fat is provided.
Introduction
Prions are the causative agents of fatal neurodegenerative
diseases, among them Creutzfeldt–Jakob disease (CJD) of man,
bovine spongiform encephalopathy (BSE) and scrapie of sheep
(see reviews by Prusiner, 1998, 1999 ; Fisher et al., 1998 ; Brown
& Bradley, 1998). They are defined as proteinaceous infectious
particles, and so far the prion protein (PrP) is the only
component that correlates unequivocally with infectivity. In
the infected organism, PrP is present both in the cellular form,
PrPC, as well as in an abnormal, scrapie isoform, PrPSc. Upon
purification using detergents and digestion with proteinase K,
PrPSc is transformed into an N-terminally truncated but still
infectious form of 27–30 kDa designated PrP 27–30 (McKinley
et al., 1991). PrPSc and the truncated PrP 27–30 form large and
insoluble aggregates. These are visible in the infected brain in
Author for correspondence : Detlev Riesner.
Fax j49 211 81 15167. e-mail riesner!biophys.uni-duesseldorf.de
† Present address : Institut fu$ r Molekulare Biotechnologie, Abt.
Molekularbiologie, Postfach 100 813, D-07708 Jena, Germany.
0001-7431 # 2001 SGM
a wide variety of sizes and shapes, i.e. from rather diffuse
depositions to solid plaques or amyloidic fibrillar forms
consisting mainly of PrP 27–30 ; the latter are called also prion
rods or scrapie-associated fibrils. Those aggregates result from
hydrophobic interactions, and indeed the prion protein has
been described as a hydrophobic protein (Prusiner et al., 1981).
The lipophilic character of the scrapie agent has been longknown, and was first inferred from its insolubility in water and
association with membranes (Hunter et al., 1971). The cellular
form of PrP, PrPC, is not present in aggregates, but is attached
to and spread on the cell membrane by a glycolipid anchor
(Naslavsky et al., 1997). In prion rods the lipids onto which PrP
was attached can still be detected (Klein et al., 1998). Thus, PrP
can be assumed to have a strong tendency for hydrophobic
interactions and to bind lipids.
Prions are characterized by an unusual resistance to the
thermal or chemical treatments which are commonly used to
inactivate agents of infectious diseases (reviewed by Casolari,
1998 ; Danner, 1991). The stability and the tendency to form
hydrophobic aggregates appear closely connected. As an
example, scrapie infectivity withstood even higher tempera-
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T. R. Appel and others
tures when heated in water in the presence of activated carbon
(Taylor, 1991). The hydrophobic surface of activated carbon
was held responsible for this effect. From studies of some
bacteria it is known that lipids effectively stabilize against
inactivation by heat (Sendhaji & Loncin, 1977 ; Sendhaji, 1977).
It is, however, not known to what extent the binding of lipids
and other components might contribute to the heat stability of
prions.
The extraordinary stability of the scrapie agent to physical
and chemical inactivation is considered today as the major
cause of the BSE epidemic resulting from the feeding of
insufficiently inactivated meat-and-bone meal to cattle (reviewed by Nathanson et al., 1997). More than 177 000 cattle in
the United Kingdom succumbed to BSE (UK Ministry of
Agriculture Fisheries and Food statistics as at 11\08\2000)
and 1 million or more infected animals may have entered the
human food chain undetected (Anderson et al., 1996). A new
variant of CJD (vCJD) affecting younger patients is most
probably caused by the consumption of infected cattle products
(Collinge et al., 1996 ; Scott et al., 1999). Measures have been
taken to reduce the potential danger of infecting humans via
cattle products, e.g. the banning of specified bovine offal and
beef on the bone from the human food chain. However, less
obvious routes of human exposure to infectious material might
also exist. Bovine tallow and bone fat are widely used as raw
material for oleochemical processes, soap and detergents,
cosmetics and animal feed. As remote as the possibility of
human infection through these products may be, it needs
further investigation when considering that more than 1n4
million metric tons of bovine fat is processed in the European
Union each year.
Almost all technical processes applied to bovine fat involve
heating, usually in the presence of water. The most prominent
example is hydrolytic fat splitting yielding free fatty acids and
glycerol. In this process temperatures exceeding 200 mC are
applied for at least 20 min in a water-saturated atmosphere
under pressure. The high potential of such treatments to
inactivate prion infectivity is obvious, but has never been
tested systematically.
This study was intended to provide quantitative data on
the heat stability of prion rods and PrP in lipid, water and
lipid–water mixtures. This is a question of general interest
because of the hydrophobic nature of prions, and also
important for assessing the safety of rendering procedures,
since several of them – as pointed out above – are performed
at high temperatures in the presence of fats and lipids. If
infectious PrP aggregates are destroyed at high temperatures
even in the presence of lipids, most products from bovine fat
could be considered safe regarding prions.
Methods
Biological safety. Prion material was handled in a BSL-2
biocontainment laboratory. Infectious samples were manipulated in
biosafety hoods using two layers of disposable gloves and oversleeves
EGG
with the upper glove discarded whenever leaving the biosafety hood.
Waste was decontaminated by autoclaving at 134 mC for 4 h or by
treatment with 1 M NaOH for 1 h at room temperature if high
temperatures could not be applied.
Prion protein samples. Recombinant prion protein rPrP(90–231)
from Syrian hamster corresponds to the sequence of prion rods PrP
27–30, but is lacking Asn-linked glycosylation and the C-terminal
glycophosphatidylinositol anchor. A sample of rPrP(90–231) was kindly
provided by Stanley B. Prusiner and Hana Serban (University of
California, San Francisco, USA). It was prepared as described previously
(Mehlhorn et al., 1996) and stored at a concentration of 10 mg\ml in
20 mM sodium acetate buffer, pH 5n5, 0n005 % thimerosal.
Prion rods (PrP 27–30) were kindly provided by Heino Diringer
(Robert-Koch-Institut, Berlin, Germany). Preparation of prion rods from
brains of terminally scrapie-sick Syrian hamsters was described earlier
(strain 263K ; Diringer et al., 1997).
PrP-specific antibody 3F4. Monoclonal antibody 3F4 (IgG2A,
kappa chain, ascites fluid, 6 mg\ml protein) was purchased from Senetek
Research, Tesson Grove Medical Center, St Louis, MO 63128, USA.
Bovine bone fat. A fat sample was provided by Henkel KGaA,
Du$ sseldorf, Germany. The fat had been recovered from bovine bones on
an industrial scale by a German processing plant. Procedures for fat
recovery consisted of squeezing the bones and mild heating.
Heat treatment of PrP in water, lipid and lipid–water
mixtures. A 300 ml pressure reactor with external electric heating,
temperature control and stirrer (Parr 4561 reactor with Parr 4842
controller) was loaded with prion rods or rPrP(90–231) in 50 ml deionized
water or in 30 g bovine bone fat or a in mixture of 50 ml water and 10 g
bone fat. The reactor was heated to the target temperature under stirring
and the start time was taken 5 mC before the target temperature was
reached. Pressure inside the reactor resembled the vapour pressure for
water at the respective temperature except in the case of pure lipid, when
less than 2 bar was observed in all cases. At the end of the reaction timeperiod, the heater was removed and the reactor cooled to 40 mC by partial
immersion in cold water.
Recovery of PrP from lipid-containing mixtures. To quantify
the amount of undegraded PrP after heat treatment in the presence of a
large excess of lipid, a method published by Wessel & Flu$ gge (1984) for
purifying proteins from a lesser amount of lipid contamination was
adapted. This method was also used for isolation of PrP after heat
treatment in pure water in order to provide results comparable to the
other experimental series involving lipids.
The contents of the reactor were transferred into a 300 ml Erlenmeyer
flask ; 300 µg BSA (bovine serum albumin fraction 5, Boehringer) was
added as a carrier protein, and the mixture stirred for 1 min. After
addition of 120 ml methanol and 60 ml chloroform for homogenization
(180 ml chloroform if 30 g lipid was present) the mixture was stirred for
2 min. Water (60 ml) was then added to separate the organic and aqueous
phases. After centrifugation (9000 g, 2 min) the proteins (BSA and PrP)
accumulated in the interphase. Most of the upper and lower phases were
removed by aspiration. The remainder contained all the protein in less
than 10 ml upper and lower phase combined. Methanol (24 ml) and
chloroform (12 ml) were added to keep all lipid in homogeneous solution
and to precipitate the proteins. The suspension was mixed thoroughly
and centrifuged (9000 g, 8 min). The proteins formed stable pellets and
the organic solvent with residual lipids could be decanted. Control
experiments with known amounts of rPrP(90–231) or PrP 27–30 showed
that more than 90 % of the protein was present in the final pellet in all
cases.
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Thermal stability of PrP
Quantification of PrP by SDS–PAGE, Western blot and
antibody detection. The method was modified from Beekes et al.
(1995) and was first published by Bendheim et al. (1984). The protein
pellets were resuspended in 500 µl loading buffer (125 mM Tris–HCl
pH 6n8, 10 % 2-mercaptoethanol, 4 % SDS, 20 % glycerol), kept for 5 min
at 105 mC and applied to SDS–polyacrylamide gels (4\12 % ; 3 mm wide).
Only every other slot was used for the analysis to avoid overlap of broad
bands. After electrophoresis the protein was transferred by electroblotting onto PVDF membrane (1 h, 2 mA\cm#, 20 V). PrP was
detected using the monoclonal antibody 3F4 (Kascsak et al., 1987), a
horseradish peroxidase-conjugated secondary antibody, followed by
enhanced chemiluminescence (Amersham ECL kit) and photographic
detection (Kodak X-ray film). The sensitivity threshold of this method
was 10 ng PrP. Developed films were digitized using a Hewlett–Packard
4c flatbed scanner with transparency adapter at a resolution of 300 d.p.i.,
256 grays, and saved as uncompressed TIFF files. PrP bands were
compared with at least three standards on the same gel using the program
Scion Image (free download from http :\\www.scioncorp.com).
A
105
0·02
0·06
0·2
E
110
10
K
L
T in °C
Fig. 2. Thermal stability of rPrP(90–231) in a lipid–water mixture. The
section of the Western blot depicted includes the interface between
stacking and resolving gels (cf. band at the top in lane B). Lanes (A)–(F),
protein pellets obtained after hydrolysis of 100 µg rPrP(90–231) in a
mixture of 10 g bovine bone fat and 50 g water for 20 min at different
temperatures (above). Lanes (G)–(L), rPrP(90–231) standards : (G)
10 ng, (H) 50 ng, (I) 100 ng, (K) 200 ng, (L) 400 ng. Right, molecular
mass in kDa.
the detection limit of 10 ng was constant. PrP bands from
samples with higher concentration appear as double bands (the
major part of the PrP with two glycosylations and a minor
portion with one glycosylation). Between 110 and 130 mC
(lanes E–G) a band of 55 to 60 kDa was detected in addition to
the main PrP band at 27–30 kDa. This corresponds to a PrP
dimer and has been described earlier (Priola et al., 1995). The
somewhat fuzzy appearance of the PrP bands, particularly if
the test lanes are compared with the standard lanes, was most
probably due to the large excess of 300 µg BSA over
0n01–0n1 µg PrP. The excess was necessary for the quantitative
precipitation of samples from the lipid–water or lipid solution,
but was used also with water-only conditions for standardization of the procedure. Every point was determined in duplicate,
showing that the conclusions drawn below were not affected
by experimental variation.
Fig. 1 shows a Western blot of prion rods heated in pure
water at different temperatures for 20 min (lanes D–K).
Quantitative values for PrP remaining undegraded were
derived by comparison with untreated controls (lanes A–C)
after densitometric evaluation. Experiments at 150 and 160 mC
(lanes I and K) were carried out with higher amounts of prion
rods compared to the experiments at lower temperatures in
order to determine the higher degradation efficiencies when
D
100
10
I
18
PrP in water
C
H
30
Known amounts of prion rods or rPrP(90–231) were added
to mixtures of bovine bone fat and water or to pure bovine
bone fat, and were heated in a laboratory autoclave to different
temperatures. Pressure inside the autoclave was similar to the
vapour pressure for water at the respective temperature except
in the case of pure lipid, when less than 2 bar was observed.
The amount of protein remaining undegraded after heating to
specific temperatures was determined according to a protocol
(see Methods) which had been developed especially for
quantitative analysis of small amounts of prion protein in the
presence of large excesses of bovine fat.
B
D
E
F G
125 130 135
60
Results
A
B
C
115 120
PrP in lipid–water mixtures
Fig. 2 presents results of experiments analysing rPrP(90–
231) heated at different temperatures in a lipid–water mixture
F
120
10
G
130
10
H
140
10
I
150
50
K
160
500
T in °C
m in µg
30
18
Fig. 1. Thermal stability of prion rods in water. Western blot visualized by PrP-specific monoclonal antibody 3F4 and enhanced
chemiluminescence detection. The slots, the stacking gel and the uppermost part of the resolving gel are not depicted ; those
parts were free of any detectable staining. It should be noted that higher amounts of PrP were used for the analysis at 150 and
160 mC in order to increase the sensitivity after extensive degradation. Lanes (A)–(C), prion rod standards. Lanes (D)–(K),
protein pellets obtained after hydrolysis of various amounts (above) of prion rods in 50 ml deionized water for 20 min at
different temperatures (above). Right, molecular mass in kDa.
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T. R. Appel and others
A
B
C
D
0·06 0·02 0·2
0·6
E
105
5
F
110
5
G
115
10
H
120
10
I
125
10
K
130
20
L
135
20
T in °C
m in µg
60
30
18
M
N
O
0·02
0·06
0·2
P
140
20
R
145
20
S
150
40
T
160
40
T in °C
m in µg
60
30
18
Fig. 3. Thermal stability of prion rods in a lipid–water mixture. Section of the Western blot depicted as in Fig. 2. Lanes (A)–(D),
(M)–(O), prion rod standards. Lanes (E)–(L), (P)–(T), protein pellets obtained after hydrolysis of various amounts (above) of
prion rods in a mixture of 10 g bovine bone fat and 50 g water for 20 min at different temperatures (above). Right, molecular
mass in kDa.
for 20 min (lanes A–F). Untreated standards were loaded for
comparison (lanes G–L). No PrP was detected at temperatures
higher than 120 mC. In addition to the monomolecular band,
aggregates of higher molecular mass were observed at 105 and
115 mC. This may be due to aggregation of PrP at these
temperatures. However, the lack of bands corresponding to
PrP dimers, trimers etc. argues for a process of random protein
splitting and aggregation of PrP with these peptides or with
components of bovine fat. At higher temperatures (lanes B and
C) much less high-molecular mass aggregate was present. The
reason might be a faster PrP splitting compared to aggregation
at higher temperatures, but the sensitivity threshold of the
antibody\chemiluminescence detection system may also play
a role.
Fig. 3 shows results of prion rod degradation experiments
at different temperatures in a lipid–water mixture. To compensate for the more effective degradation at higher temperatures the amount of prion rods used was increased with
temperature (see lane headings of Fig. 3). Thus, the effect of
temperature is much more pronounced as seen directly from
the Western blot. In the experiments with prion rods as
compared to rPrP(90–231), PrP was still detectable at much
higher temperatures, i.e. 160 mC instead of 120 mC for 20 min.
It should be noted that signals at 60 kDa indicative of PrP
dimers are seen in Fig. 3, lanes (E)–(L). These dimers may have
formed de novo as a result of high temperature treatment but
could also have already been present in prion rods.
EGI
A
0·06
B
130
3
C
140
7·5
D
150
37·5
E
160
375
T in °C
m in µg
200
60
30
18
Fig. 4. Thermal stability of prion rods in lipid. Section of the Western blot
depicted as in Fig. 2. Lane A, prion rod standard. Lanes (B)–(E), protein
pellets obtained after hydrolysis of various amounts (above) of prion rods
in 30 g bovine bone fat for 20 min at different temperatures (above).
Right, molecular mass in kDa.
PrP in pure lipid
PrP inactivation experiments using water or a lipid–water
mixture at different temperatures (Figs. 1 to 3) displayed the
main PrP band at 27–30 kDa. Densitometric evaluation of
these experiments was possible by comparison to PrP
standards on the same gel. In contrast, when heating prion rods
in bovine bone fat in the absence of water, PrP signals were
observed exclusively at higher molecular masses (Fig. 4). At
130 mC (lane B) bands indicative of PrP dimers, tetramers or
octamers were seen at about 60, 120 and 240 kDa. At higher
temperatures (lanes C–E) a signal was detected only at the
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Thermal stability of PrP
Table 1. Logarithmic values of the degradation factors
for prion rods and rPrP(90–231) in water or lipid–water
mixtures for 20 min at different temperatures (DF20)
log10 (mstart/mend)
Prion rods in
H2O
Prion rods in
H2O–lipid
rPrP(90–231) in
H2O–lipid
100
105
110
115
1n8

2n5
2n8

0n7

2n0
120
2n8
3n0


0n5
0n6
0n9
1n0
1n2
1n3
1n3
1n4
1n8
1n8
1n8
1n9
1n8
2n2
T (mC)
125
130
135
140
2n7
3n1

160
3n0
3n1
3n9

4n2
4n6
4n8
170

145
150
3n0
2n9
3n0
3n6
3n7
4n0
3n0
4n0







PrP aggregates withstood boiling in sample buffer containing
SDS and 2-mercaptoethanol. Besides the formation of stable
aggregates (as seen in Fig. 4) decomposition of PrP occurred at
high temperatures in bovine bone fat. Quantitative values
cannot be given because of the lack of standards for PrP
aggregates, but from the known sensitivity threshold of 10 ng
and the amounts of prion rods needed to give a signal after
heating to 160 mC (Fig. 4, lane E) a reduction factor of about
10% can be assumed at this temperature.
Quantitative evaluation
The results of the densitometric analyses of Figs 1, 2 and 3
as well as further series of prion rods in water or a lipid–water
mixture at different temperatures (not shown) are presented in
Table 1. The logarithmic degradation factors (DF) are listed, i.e.
the ratio of the starting amount of PrP divided by the amount
remaining undegraded after autoclaving for 20 min at the
respective temperature.
The heat inactivation kinetics of prions can be described by
a first-order decay reaction (Casolari, 1998 ; Overthu$ r, 2000) :
CPrP(t) l C!PrP×e−kt
(1)
! the concentration of undegraded PrP at the start,
with CPrP
CPrP(t) the concentration of undegraded PrP after the inactivation time t and k the rate of decay.
The degradation factor (DF) (see above) was defined as :
DF l C!PrP\CPrP(t)
leading to
, Not determined.
interface between the stacking and the resolving gel. This
indicates an aggregation process of PrP yielding very stable
aggregates with a molecular mass higher than 350 kDa. These
log DF l kit\2n303
"!
For an inactivation time of 20 min :
(2)
log DF l ki1200\2n303
"! #!
(3)
Fig. 5. Logarithmic degradation factors
(DF20) obtained after heating prion rods
or rPrP(90–231) in water or lipid–water
mixtures for 20 min. The data are plotted
in the form of an Arrhenius plot according
to eqns (3) and (4) with linear
regression. $ (——), Prion rods in
water ; (– – –), prion rods in
water–lipid ; > (:::), rPrP(90–231) in
water–lipid.
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T. R. Appel and others
Thus, the log DF values are proportional to the rate
"! #!
constants of the inactivation reaction at the respective
temperature.
The temperature dependence of rate constants is given by
the Arrhenius equation :
(4)
clnk\c1\T lkEa\R
with EA being the activation energy for the inactivation
process and R the gas constant.
Because log DF is proportional to the rate constant k, a
"! #!
plot of ln(log DF ) versus 1\T is presented in Fig. 5. A linear
"! #!
relationship can be seen for all series of prion rods or
rPrP(90–231) in water or lipid–water mixtures.
Discussion
Methodological aspects
PrP is the major component of prion rods, which are the
purest preparation of a prion-type agent. Prion rods consist
mainly of the N-terminally truncated form PrP 27–30, which is
highly aggregated and has a secondary structure rich in βsheet. In addition to PrP 27–30, prion rods also contain about
1 % (m\m) sphingolipids (Klein et al., 1998) and 10–15 %
(m\m) of an inert polysaccharide scaffold (Appel et al., 1999).
If prion rods are heated, non-covalent interactions and, at
higher temperatures, chemical bonds are destroyed at an
increasing rate. Aggregates can form or dissociate into
monomers or oligomers of PrP, tertiary and secondary
structures of PrP denature and peptide bonds may break.
Under the conditions of the experiments in this work watermediated hydrolysis of the peptide bond most probably
represents the major contribution to the degradation observed.
The hydrolytic attack of water on the peptide bond is expected
to increase proportional with temperature.
The degree of inactivation of Syrian hamster prion rods
was determined by the amount of PrP appearing in Western
blots as an undegraded monomeric band or as high molecular
mass aggregates after heat treatment. Since monoclonal
antibody 3F4 recognizes the sequence of PrP amino acids
109–112, this sequence has to be undegraded in the monomers
as well as in the high molecular mass aggregates. Only the
amount of monomers could be evaluated quantitatively and
was the basis for the quantitative conclusions.
Prion infectivity diminishes concomitantly with PrP proteolysis (Prusiner, 1982). All reports published so far show that
prion infectivity is proportional to PrP (McKinley et al.,
1983 ; Gabizon et al., 1989 ; Brown et al., 1990 ; Beekes et al.,
1996). In conclusion, if a chemical or physical treatment
destroys the peptide bonds of PrP, it also destroys prion
infectivity. Prion infectivity might, however, also be degraded
by denaturing mechanisms other than degradation (cf. above) ;
consequently degradation factors, DF, calculated from the
amount of undegraded PrP are minimum factors for inactivation of infectivity. One should note that the loss of PrP
in the 27–30 kDa band as evaluated in the present work could
EHA
also be due to aggregation without loss of infectivity.
Aggregation was indeed observed as a clearly visible band at
the border between stacking and resolving gels after heat
treatment in pure lipid (cf. Fig. 4), whereas after treatment in
water or a lipid–water mixture (cf. Figs 2 and 3) the amount of
aggregates in the Western blot analysis was negligible and did
not affect the analysis. Therefore, quantitative values for
degradation factors could be given only in the case of heat
treatment in water or lipid–water mixtures, but not in pure
lipid. Furthermore, because prion rods result from purification
by extended proteinase K digestion, they represent the most
stable fraction of the original infectious material (Prusiner et al.,
1983 ; Diringer et al., 1997), underlining the conclusion that the
degradation factors are very cautious estimates of the efficacy
of inactivation of infectivity.
In several aspects the Western blot test for PrP has
advantages compared to a direct measurement of scrapie
infectivity. First, infectivity cannot be easily concentrated at
the end of an inactivation process because almost any method
of concentrating PrP deactivates infectivity. Since animal
bioassays are limited by the small volume (30 µl) that can be
injected intracerebrally, only a minuscule fraction could be
tested. Thus, only moderate infectivity would be present per
sample even if highly infectious spiking material is employed
(Taylor et al., 1995, 1997). This dilution is avoided by testing
for PrP, which can be recovered almost completely from large
volumes by precipitation together with the carrier protein
BSA. Second, PrP degradation factors up to 10& can be
determined within hours by a Western blot assay, whereas
bioassays need 200 days or more for completion. Third, the
relation between incubation time and dose of at least some
strains of scrapie is altered upon heating (Taylor et al., 1996).
Therefore expensive and time-consuming end-point dilution
assays would have to be used for testing prion infectivity after
heating.
It was not possible to measure protein recovery by a
second independent method, e.g. amino acid analysis of the
chloroform–methanol precipitate. The large excess of BSA
needed for quantitative co-precipitation of PrP and residual
bovine proteins present in the technical bovine bone-fat
excluded any method other then the specific PrP staining by
antibodies.
Mechanistic implications for the protective effect of
lipids
The results summarized in Fig. 5 show that the presence of
lipids increases the heat stability of prion rods (dashed line vs
solid line). This difference is more than two orders at lower
temperatures ($ 100 mC), but decreases with increasing temperature and extrapolates to about the same stability between
160 and 170 mC. One reason for the observed protective effect
of lipids on prion rods may be additional hydrophobic
interactions. Lipids can saturate the hydrophobic surface of PrP
and protect it from hydrolytic attack by replacing the water
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Thermal stability of PrP
layer surrounding prion rods. Conventional infectious agents
like bacteria are also protected against heat by lipids (Sendhaji & Loncin, 1977 ; Sendhaji, 1977), but at a much lower
temperature range.
As mentioned, the protective effect of lipids decreases with
increasing temperature. Two alternative explanations could be
considered. One would be that the non-covalent associations
with lipids that protect prion rods against hydrolytic attack are
gradually lost with increasing temperature. Thus, at higher
temperatures ‘ regular ’ hydrolytic processes resume. In that
case the degradation curves of prions in lipid–water mixtures
and in pure water should be extrapolated to higher temperatures according to the curve in pure water. An alternative
scenario assumes that random bond-breaking due to Brownian
motion is the dominant inactivation effect for PrP–lipid
complexes because of the protection against hydrolytic attack.
In that case inactivation of prion rods in both pure water and
the lipid–water mixtures has to be extrapolated to higher
temperatures according to the curve in lipid–water mixtures.
Unfortunately, these alternatives cannot be differentiated at
present because of the limited sensitivity of PrP detection,
which did not allow us to analyse the PrP remaining at
temperatures above 160 mC.
A somewhat different effect was observed with prion rods
heated in pure bovine bone fat at high temperatures (Fig. 4). In
contrast to water or lipid–water mixtures, no monomeric PrP
band was obtained. Instead, bands indicative of dimers,
tetramers and higher aggregates were present. The aggregates
were formed irreversibly, i.e. were retained in the aqueous and
denaturing conditions of the gel electrophoresis before the
Western blot, implying covalent links between the PrP
molecules. Covalent links between individual PrP molecules
might be formed at elevated temperatures by reactions starting
from the double bonds that are present abundantly in lipids or
from Maillard reactions between free side-chain amino groups
of PrP and carbohydrates, i.e. from the polysaccharide scaffold
of prion rods, the Asn-linked glycosylations or the glycophosphatidylinositol anchor of PrP. Since inactivation by hydrolytic
attack is excluded here due to the absence of water, a similar
inactivation mechanism as in lipid–water mixtures could be
expected.
Comparison with infectivity data known from the
literature
Unfortunately, it is technically impossible to carry out
prion infectivity studies in the lipid–water mixtures or lipids as
used in this study. As outlined above the sensitivity would be
too low to follow inactivation over a larger temperature range.
But how can the results on the heat stability data for prion
protein derived from PrP Western blot assays be compared to
literature data obtained from measuring infectivity ? Only
limited comparisons are possible because almost every former
study used different scrapie material, like brain homogenate,
various extracts thereof, partially purified rods etc. at different
Table 2. Logarithmic values of the degradation factors
for 263K prion rods or brain homogenate at different
temperatures and exposure times
Comparison of biochemical data from this study (bold type) with
infectivity data from the literature.
log10 (mstart/mend)
Prion rods
T (mC)
t (min) In H2O
120
121
20
60
130
132
90
20
60
134
135
140
90
18
30
20
20
2n9
In H2O–lipid
Brain homogenate
10 %
100 %
1n2
7n6*
8n3†
(5n3)‡
(5n6)‡
2n9
1n8
3n2§
3n5
8n8‡
(6n7)‡
( 7n4)‡
2n1R
2n0R
1n9
2n0
* Brown et al. (1982) (in PBS).
† Brown et al. (1986) (in PBS).
‡ Ernst & Race (1993) (in PBSj0n32 M sucrose).
§ Brown et al. (1990).
R Taylor et al. (1994).
temperatures and exposure times. Systematic compilations of
these data have been published by Overthu$ r (2000), Casolari
(1998) and Danner (1991).
Table 2 shows all available data on the heat stability of the
hamster-adapted scrapie strain 263K used in this study. Brown
et al. (1990) reported a decrease of 3n2 log ID for purified
&!
263K prion rods in PBS at 134 mC for 30 min. This value agrees
very well with our data on prion rods in water as quantified by
Western blot. Other results found in the literature were all
obtained by heating homogenates of terminal 263K scrapieinfected hamster brain, either undiluted (100 %) or diluted
tenfold by PBS or PBS containing 0n32 M sucrose (10 %).
Inactivation factors obtained from tenfold-diluted brain homogenates were orders of magnitude higher than ours. As
outlined above a difference could be expected from the
structural difference of prions in brain tissue and prion rods
after preparation. It is known that PrPSc in the brain has a more
diffuse structure, which is more accessible to water hydrolysis
than the extremely compact prion rods. Furthermore, the high
inactivation factors obtained with 10 % homogenate do not
contradict the data of the present work obtained with prions in
pure water, if the exponential decay with time were to be
extrapolated from 20 min to 60 or 90 min, respectively. On
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T. R. Appel and others
Table 3. Degradation factors (DF20) for 263K prion rods in water or lipid–water mixtures at different temperatures
Temperature (mC)
100
DF in water
#!
DF in lipid–water
#!
10#
10"
110
120
130
140
150
160
170
180
190
200
3i10#
10"
6i10#
10"
10$
3i10"
3i10$
10#
10%
6i10#
2i10%
5i10$
10&
10&
(4i10&)*
(4i10&)
(2i10')
(2i10')
(10()
(10()
* Values in parentheses are extrapolated from Fig. 5.
the other hand, the data of Taylor et al. (1994) obtained with
undiluted brain homogenate at 134 mC are almost identical to
our results determined in a lipid–water mixture. This is most
probably due to the large amounts of lipids present in undiluted
brain tissue.
In summary, the degradation factors derived from Western
blots of PrP in this work were similar to the inactivation factors
from infectivity tests, if data obtained under equivalent
conditions were compared. The compilation of data in Table 2
demonstrates the critical effect that the solvent conditions
have on the efficiency of the inactivation process.
The authors are indebted to Dr Stanley B. Prusiner and Ingrid
Mehlhorn (San Francisco, USA) for a kind gift of rPrP(90–231) and to Dr
Heino Diringer (Berlin, Germany) for a kind gift of prion rods. Many
thanks to Radulf C. Overthu$ r for helpful discussions and a preprint of his
manuscript on the heat inactivation of prions. The work was supported
by grants from the Bundesministerium fu$ r Bildung und Forschung (FRG).
T. R. A. is a fellow of the Du$ sseldorf entrepreneurs foundation.
Conclusions
Anderson, R. M., Donnelly, C. A., Ferguson, N. M., Woolhouse, M. E.,
Watt, C. J., Udy, H. J., MaWhinney, S., Dunstan, S. P., Southwood,
T. R., Wilesmith, J. W., Ryan, J. B., Hoinville, L. J., Hillerton, J. E., Austin,
A. R. & Wells, G. A. (1996). Transmission dynamics and epidemiology of
Regulatory publications have assumed a prion inactivation
factor of 10) for autoclaving at 133 mC for 20 min [Bundesministerium fu$ r Gesundheit (1996) ; similar values given by UK
Department of Health and Social Services (1984) and American
Neurological Association (1986)]. This value appears rather
high in view of the present results, even if one considers that
it is applied mostly for dilute solutions in the absence of excess
lipids, as holds true in most medical or technical applications.
Furthermore, it should be noted that the experimental data
underlying the regulatory prescriptions have been derived
from experiments with mouse scrapie or BSE, which are less
stable to autoclaving than the experimental 263K hamster
scrapie strain used here (Taylor et al., 1994). However, 100 %
brain homogenate or mixtures containing large amounts of
lipids may behave differently. Under these circumstances, our
comprehensive data on the heat inactivation of 263K hamster
prion rods in water or a lipid–water mixture represent
minimum inactivation limits for the temperature range 100 to
170 mC. When extrapolating Fig. 5 to temperatures above
170 mC, only the lower slope of the curve for prion rods (solid
line) in water should be used. Although the theoretical aspects
of the alternatives for extrapolation have been discussed
above, lack of experimental data and safety considerations
allow us only to recommend the lower slope for extrapolation
above 170 mC ; i.e. that of prions in pure water. Extrapolation
to 200 mC yields a degradation factor of 10(, representing a
fairly good safety limit. Table 3 gives an overview of
degradation factors from Fig. 5 and their extrapolation to
200 mC.
EHC
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