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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- Downloaded from www.microbiologyresearch.org by IP: 88.99.165.207 On: Sun, 18 Jun 2017 23:49:11 EGF 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. Downloaded from www.microbiologyresearch.org by IP: 88.99.165.207 On: Sun, 18 Jun 2017 23:49:11 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. Downloaded from www.microbiologyresearch.org by IP: 88.99.165.207 On: Sun, 18 Jun 2017 23:49:11 EGH 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 Downloaded from www.microbiologyresearch.org by IP: 88.99.165.207 On: Sun, 18 Jun 2017 23:49:11 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. Downloaded from www.microbiologyresearch.org by IP: 88.99.165.207 On: Sun, 18 Jun 2017 23:49:11 EGJ 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 Downloaded from www.microbiologyresearch.org by IP: 88.99.165.207 On: Sun, 18 Jun 2017 23:49:11 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 Downloaded from www.microbiologyresearch.org by IP: 88.99.165.207 On: Sun, 18 Jun 2017 23:49:11 EHB 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. 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