The centromeric URA3 plasmid pRS316 (53) and 2μm DNA-based multicopy plasmids YEp13, bearing the LEU2 marker (4), and pEMBL-yex, bearing the URA3 and LEU2-d markers (5), were described previously. Plasmid pRS316GAL is a pRS316 derivative bearing the galactose-inducible (GAL) promoter GAL1,10 (37). The centromeric plasmids pLA1, which contains the HIS3 marker and GAL promoter, and pH28, which is a pLA1 derivative bearing the HSP104 gene control of under the GAL promoter, were described previously (39). The centromeric plasmid pYS-GAL104 (11, 35) contains the HSP104 gene under control of the GAL promoter and the URA3 marker. The centromeric LEU2 plasmid pLH105, kindly provided by S. Lindquist, contains the HSP104 gene under control of the constitutively expressed GPD promoter. The previously described plasmids pEMBL-SUP35 (also called pEMBL-SUP2) and pEMBL-SUP35-ΔBal (also called pEMBL-SUP2-ΔBal) are pEMBL-yex derivatives which bear the complete S. cerevisiae SUP35 gene and the N-terminal 154 codons of the SUP35 gene, respectively, under control of the endogenous SUP35 promoter (55). The centromeric URA3 plasmid CEN-GAL-SUP35 (also called pVK71) contains the complete S. cerevisiae SUP35 gene under control of the GAL promoter (21). Plasmid pRS316GAL-SUP35Pm (12) is a pRS316GAL derivative which contains the Pichia methanolica SUP35 open reading frame (ORF) (32) placed under the control of the GAL promoter. The centromeric URA3 plasmid YCp-SSB1, which is a pRS316 derivative bearing the wild-type SSB1 gene under control of its endogenous promoter, was kindly provided by M. Ohba (41). Plasmids pRS316GAL-SSB1#1 and pRS316GAL-SSB1#2 were constructed by PCR amplifying the SSB1 ORF from the genome of S. cerevisiae S288C and placing it into pRS316GAL under control of the GAL promoter. Primers SSB1.EXT5 (5′-TACAGGATCCGTCCCAAGATCATTAC-AGTATT) and SSB1.EXT3NEW (5′-TACAGCGGCCGCCATATATATGTGATGAATGCAG), which are complementary to the 5′ flanking region (positions −35 to −14) and 3′ flanking region (positions +1845 to +1866) of the S. cerevisiae SSB1 ORF, respectively, were used in PCR with Pfu polymerase. These primers contain extensions bearing the restriction sites for BamHI and NotI, correspondingly. The PCR products were cut with BamHI and NotI and ligated into the BamHI-NotI cut pRS316GAL. To minimize the possible effect of PCR-generated errors, two independent PCRs were performed, each leading to one plasmid construct (1 or 2). Resulting plasmids pRS316GAL-SSB#1 and pRS316GAL-SSB#2 produced identical results in all experiments described below and are further designated pRS316GAL-SSB1. On galactose medium, either plasmid compensated for the ssb1/2Δ-associated phenotypes, such as paromomycin sensitivity (Par^s) (38) and GuHCl sensitivity (Ghc^s) (see below). Western blot results confirm that yeast strains bearing any of these plasmids exhibit increased levels of the Ssb protein on galactose medium (40). Plasmids pBS-HIS3-I and pBS-URA3-I, used in gene disruption experiments (see below), were constructed by inserting the 1.7-kb fragment bearing the complete HIS3 gene and the 1.1-kb fragment bearing the complete URA3 gene, respectively, into pBluescript KS II(+) polylinker. The URA3 centromeric plasmids pUKC815 and pUKC819, which were used for the quantitative measurement of UGA suppression, were kindly provided by M. F. Tuite. These plasmids contain the in-frame fusions PGK-lacZ and PGK-UGA-lacZ, respectively (25).

The direct PCR-mediated transplacement protocol (2) was used to disrupt SSB1 and SSB2 genes. Primers SSB1-BLUE.PRO (5′-GATGT CCCAAGATCATTACAGTATTTTAATTGAACCTCACTATAGGGCGAA) and SSB1-BLUE.TER (5′-TAAGTAATATTCATATATATGTGATGAATGCAGTCCCTCACTAAAGGGAACAA) were designed in such a way that their 5′ flanking portions were complementary to the 5′ and 3′ flanks of the chromosomal SSB1 gene, respectively. The 3′ portions of the same primers were complementary to sequences of the pBluescript KS II(+) polylinker, located on the 5′ and 3′ sides, respectively, of the HIS3 insertion in plasmid pBS-HIS3-I. These primers were used to PCR amplify the HIS3 fragment from pBS-HIS3-I in a Taq polymerase-mediated reaction. The resulting HIS3 fragment, which contains the complete HIS3 gene flanked by extensions homologous to the flanks of the chromosomal copy of SSB1, was gel purified and transformed into strain GT81-1C. Transformants were selected on His-deficient (−His) medium. The ssb1Δ::HIS3 disruptants, in which the complete SSB1 ORF is deleted and replaced by the HIS3 gene in the same orientation, were verified by Southern hybridization as described below. One of these disruptants, GT128, was used in further transplacement experiments employing primers SSB2-BLUE.PRO (5′-TTTCAAGA AACCAAGAACCAATATCCTCATTAACACTCACTATAGGGCGAATT) and SSB2-BLUE.TER (5′-ATATATATGTGTATAACCTTAACCAGAATGACATCCCTCACTAAAGGGAACAA) to amplify the URA3 gene from plasmid pBS-URA3-I. The resulting URA3 fragment, flanked by extensions homologous to the 5′ and 3′ flanks of the chromosomal SSB2 gene in the same orientation, was gel purified and transformed into strain GT128. Transformants were selected on −Ura medium. One, containing a double ssb1Δ::HIS3 ssb2Δ::URA3 (ssb1/2Δ) disruption, was verified by Southern hybridization as described below. This ssb1/2Δ strain and its derivatives exhibited slow growth at low temperature (20 to 25°C) and sensitivity to the aminoglycoside antibiotics paromomycin and hygromycin, as described previously (38).

Standard yeast media, cultivation conditions, and standard procedures for yeast growth, sporulation, micromanipulation, and tetrad analysis were used (30). Gal medium contained 2% galactose instead of glucose. Gal+Raf medium contained 2% galactose and 2% raffinose instead of glucose. Transformation was performed according to a modified Li⁺ protocol (30). Yeast cultures were grown at 30°C unless specified otherwise. Liquid cultures were grown on a shaker, normally at 200 to 250 rpm, with a liquid/flask volume ratio of 1:5 or more.

To assay for [PSI] curing by the overproduced Hsp104, yeast cultures bearing the GAL-HSP104 construct or matching controls were grown to 2 × 10⁶ to 1 × 10⁷ cells/ml in liquid synthetic glucose medium selective for the plasmid(s), washed twice with H₂O, and inoculated into the corresponding synthetic Gal+Raf medium at the starting concentration of 2.5 × 10⁵ to 5 × 10⁵ cells/ml. Aliquots were taken before and after incubation (usually 22 to 24 h) and plated onto the solid synthetic glucose medium selective for the plasmid(s). The colonies grown after 4 to 5 days were velveteen replica plated onto YPD and −Ade medium. [PSI⁺] colonies were identified by white color on YPD and growth on −Ade medium after 4 to 6 days of incubation.

The plate assays for [PSI] induction were performed as described previously (21). Yeast transformants bearing plasmid CEN-GAL-SUP35 or pRS316GAL-SUP35Pm, or matching control plasmid pRS316GAL, were grown on glucose −Ura medium and velveteen replica plated onto −Ura and −Ura/Gal media. After 3 to 4 days of incubation, each plate was velveteen replica plated onto glucose −Ade medium. [PSI] induction was detected as heterogeneous growth on −Ade medium after 7 to 10 days of incubation. To assay [PSI] induction quantitatively, transformants bearing the CEN-GAL-SUP35 plasmid were grown in the synthetic glucose −Ura medium, washed twice with H₂O, and inoculated into the −Ura/Gal+Raf medium at the starting concentration 10⁵ cells/ml. The aliquots were taken before incubation and after various periods of incubation and plated onto the glucose −Ura medium. The colonies grown after 4 to 5 days were velveteen replica plated onto the YPD and −Ade media. [PSI⁺] colonies were identified by white color on YPD medium and growth on −Ade medium after 6 to 8 days of incubation.

To measure the frequencies of the spontaneous [PSI] formation, yeast strains were streaked out on YPD medium at 30°C to obtain single colonies originating from individual cells; 11 to 12 colonies were analyzed for each strain. The whole colony was resuspended in 200 μl of H₂O; 10 μl of each solution was taken to perform serial dilutions in H₂O, and the rest of each solution was plated onto −Ade medium. A 1/10 aliquot of the first (1/10) dilution was taken for further serial dilutions, while the rest of the 1/10 dilution was also plated on −Ade medium. Further serial dilutions were plated on YPD medium in order to determine the concentrations of viable cells (CFU). Ade⁺ colonies were counted on −Ade plates following 10 days of incubation at 30°C. Numbers of Ade⁺ colonies in undiluted cultures and 1/10 dilutions were roughly proportional to the degree of dilution, suggesting that residual growth on −Ade medium is minimal and has no significant effect on the frequency of Ade⁺ derivatives. For each sample, the numbers of Ade⁺ colonies detected in the undiluted culture and in 1/10 dilution were summed to determine the total number of Ade⁺ derivatives in the sample. Median frequencies of the Ade⁺ colonies were calculated for each strain and shown in Table ​Table2.2. [PSI] formation rates were calculated according to the formula R = f/ln(NR), where R is rate of [PSI] formation, f is the observed frequency of [PSI⁺] colonies, and N is number of cells in the culture. This approach is based on the formula used previously for measuring spontaneous mutation rates (24).

Standard protocols were used for DNA isolation, restriction digestion, ligation, and Escherichia coli transformation (49). Oligonucleotides were purchased from Gibco BRL. Restriction endonucleases were purchased from New England Biolabs and Gibco BRL. Polymerases were purchased from Gibco BRL, Promega, and Stratagene. The thermal cycler was from Ericomp. Southern hybridization was with a chemiluminescent probe as described in the Amersham protocol. The 1.9-kb BamHI-SacII fragment of plasmid pRS316GAL-SSB1, labeled by the random priming procedure, was used as a probe. It hybridizes to both SSB1 and SSB2 genes due to high homology between their ORFs. The chromosome DNA was cut with SacII to produce a 2.8-kb fragment corresponding to the SSB1 gene and large (>20-kb) fragment corresponding to the SSB2 gene. These bands were both absent in the double ssb1Δ::HIS3 ssb2Δ::URA3 disruptant. Additional digests with XbaI and with XbaI plus SacII were also analyzed to confirm disruptions. The blot was also probed with the labeled HSP104 gene used as a DNA loading control. Proteins were isolated as described previously (39) and subjected to sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). Antibodies specific to the Hsp104, Hsp26, Hsp82, and Sup35 proteins were kindly provided by S. Lindquist. Antibodies specific to Ssa protein were kindly provided by E. Craig. Reactions with antibodies specific to Ssb protein were performed by M. Patino in S. Lindquist’s lab. Reactions with antibodies were detected by using the ECL (enhanced chemiluminescence) detection system according to the Amersham protocol. Relative amounts of proteins were determined by densitometry where necessary by using the Image Tool program as described previously (39). The amounts of proteins and times of exposure were kept within the linear range of the ECL detection system, determined on the basis of our previous data.

To check whether the absence of Ssb protein would affect [PSI], we disrupted SSB1 and SSB2 genes in the [PSI⁺] strain GT81-1C as described in Materials and Methods. Resulting Ssb⁻ disruptants remained [PSI⁺], and [PSI] remained stable in mitotic divisions. Moreover, [PSI] curing by both the GAL-HSP104 construct, induced in the Gal+Raf medium (Fig. ​(Fig.1B),1B), and the constitutively expressed GPD-HSP104 construct (Fig. ​(Fig.1C)1C) were less efficient in the Ssb⁻ strain than in the isogenic Ssb⁺ strain. Neither parameters of growth in Gal+Raf medium (Fig. ​(Fig.1B)1B) nor levels of the Hsp104 protein (Fig. ​(Fig.1E1E and F) were significantly affected, suggesting that the absence of Ssb influences [PSI] curing by Hsp104 rather than the number of generations or Hsp104 expression. This result confirms that Ssb protein assists in [PSI] curing by excess Hsp104 protein.

Inactivation of the Hsp104 protein has been shown to cure yeast cells of [PSI] (11). We checked whether [PSI] loss, caused by Hsp104 inactivation, requires Ssb protein. The [PSI⁺] ssb1/2Δ strain was crossed to the isogenic hsp104Δ strain GT84-5A. The resulting diploid was sporulated and dissected. Among 19 complete tetrads analyzed, we identified 2 which contained only two Ura⁺ spores each. In such tetrads, the ssb2Δ::URA3 and hsp104Δ::URA3 disruptions had to segregate together. The ssb1Δ::HIS3 disruption was monitored by the His⁺ phenotype, and the double ssb1/2Δ disruptants were also verified by the Par^s phenotype. Using these approaches, we determined that three of four Ura⁺ spores recovered from these two tetrads contained all three disruptions simultaneously (ssb1Δ::HIS3 ssb2Δ::URA3 hsp104Δ::URA3). One of these spore clones was also tested by Southern hybridization to confirm the presence of all three disruptions. All three triple disruptant spores were [psi⁻], confirming that [PSI] curing by Hsp104 inactivation does not require the presence of Ssb protein.

Spontaneous appearance of [PSI] is easier to detect in strains containing the non-Mendelian determinant [PIN], which increases the frequency of [PSI] formation (22). We observed that the [psi⁻ PIN⁺] Ssb⁻ strain exhibited 1-order-of-magnitude-higher frequency and rate of spontaneous Ade⁺ revertants compared to the isogenic [psi⁻ PIN⁺] Ssb⁺ strain (Fig. ​(Fig.2A2A and Table ​Table2).2). In all 12 independently arisen spontaneous Ssb⁻ Ade⁺ colonies tested, the Ade⁺ phenotype was cured by growth in the presence of GuHCl, as described previously for the [PSI] element (57). This finding suggests that the increased frequency of the Ade⁺ revertants in the Ssb⁻ strain is due to increased spontaneous appearance of the [PSI] prion.

Overproduction of the Sup35 protein causes translational readthrough, or nonsense suppression (9), and induces formation of [PSI] prion (10) in [psi⁻ PIN⁺] strains (22) of S. cerevisiae. Overproduction of the Sup35N prion-forming domain alone also induces nonsense suppression (55) and [PSI] formation (21) in both [psi⁻ PIN⁺] and [psi⁻ pin⁻] strains (22). We have observed that nonsense suppression by overproduced Sup35 or Sup35N in the [psi⁻ PIN⁺] background (Fig. ​(Fig.2B)2B) is greater in the Ssb⁻ strain than in the isogenic Ssb⁺ strain. Nonsense suppression by overproduced Sup35N in the [psi⁻ pin⁻] background was also increased in the Ssb⁻ strain (not shown), indicating that the Ssb effect is not [PIN] specific. The frequency of [PSI] formation, induced by the transient Sup35 overproduction in the [psi⁻ PIN⁺] background, was also increased 6- to 10-fold in the strain lacking Ssb protein (Fig. ​(Fig.2C2C and D). The growth rate of the [psi⁻ PIN⁺] Sup35 overproducer was only slightly influenced by ssb1/2Δ (Fig. ​(Fig.2E),2E), which was not sufficient to explain a difference in [PSI] induction rates.

The sequence variation between the prion proteins of different origins decreases the efficiency of prion conversion in the heterologous system, which is recognized as a species barrier in prion transmission (for a review, see reference 48). We have shown that the highly divergent Sup35NM domain, originated from the distantly related yeast species Pichia methanolica (32), can form a prion in the S. cerevisiae cellular environment (12). Moreover, overproduction of Pichia Sup35 induces formation of the S. cerevisiae [PSI] prion; however, the efficiency of such heterologous prion induction is much lower than that of the homologous induction by overproduction of the endogenous S. cerevisiae Sup35 (12). Our data (Fig. ​(Fig.2C)2C) indicate that the heterologous [PSI] induction by P. methanolica Sup35 is increased in the Ssb⁻ background. Therefore, Ssb depletion makes Sup35 more sensitive to the heterologous prion-inducing agents and helps to overcome the species barrier.

One possible explanation for the increased [PSI] appearance in the Ssb⁻ background could be that Ssb, which is a ribosome-associated protein (38, 47), antagonizes the translational readthrough, caused by [PSI], or prevents folding and/or facilitates destruction of the readthrough products, which likely contain the wrong amino acids inserted. In this case, the efficiency of nonsense suppression would be increased in the absence of Ssb. Therefore, some weak [PSI⁺] derivatives, which are not detectable in an Ssb⁺ background, could become detectable in an Ssb⁻ background. Indeed, in some experiments the Ssb⁻ derivative of the [PSI⁺] strain GT81-1C grew a little better than the original Ssb⁺ strain on −Ade medium, although this difference was difficult to reproduce (not shown). To check it quantitatively, we measured nonsense suppressor efficiencies by using the PGK-UGA-lacZ construct. Our results show that while the effect of ssb1/2Δ on UGA suppression in the [psi⁻] strain was insignificant, the level of UGA suppression by [PSI] was increased about fivefold in the Ssb⁻ background (Table ​(Table3).3). To check whether such an effect could compromise the [PSI] formation assays, we transformed three independent spontaneous [PSI⁺] derivatives of the [psi⁻ PIN⁺] ssb1/2Δ strain and 11 independent [PSI⁺] derivatives induced by Sup35 overproduction in the same strain with plasmid YCp-SSB1, bearing the wild-type SSB1 gene. Our results (e.g., Fig. ​Fig.2F)2F) confirm that all [PSI⁺] derivatives obtained in the absence of Ssb remain able to grow on −Ade medium in the presence of Ssb. Therefore, while Ssb protein appears to affect nonsense suppression by [PSI], results of the [PSI] formation assays cannot be explained by the Ssb effect on nonsense suppression. Moreover, reintroduction of SSB1 caused a detectable decrease in growth on −Ade medium only in 3 of 14 independent [PSI⁺] derivatives tested. We previously reported that various strains of the [PSI] prion could be recovered in the same genotypic background, and so differences between these strains are apparently controlled by the prion protein itself (21). Thus, the Ssb protein appears to have differential effects on the different prion strains.

Effects of Ssb depletion on suppression and [PSI] formation are not readily explained by alterations of the Sup35 protein levels or its distribution between soluble and precipitated (that is, aggregated) fractions, since neither of these parameters appears to be affected in the [psi⁻ PIN⁺] cells lacking Ssb (Fig. ​(Fig.3A3A to C). To determine whether ssb1/2Δ acts on [PSI] indirectly by altering other chaperone levels, we compared levels of several chaperone proteins in the isogenic Ssb⁺ and Ssb⁻ strains. Our results confirm that levels of Hsp104 protein on glucose medium are unaffected by ssb1/2Δ (Fig. ​(Fig.3D),3D), as also shown above for Gal+Raf medium (Fig. ​(Fig.1E1E and F). Levels of Ssa protein were also unaffected (Fig. ​(Fig.3D).3D). There appeared to be a slight increase of Hsp82 levels in the Ssb⁻ background (Fig. ​(Fig.3D);3D); however, it has previously been shown that [PSI] is not affected by Hsp82 levels (39). Neither Ssb⁺ nor Ssb⁻ cells produced Hsp26 protein in the exponential phase, independently of whether they were [PSI⁺] or [psi⁻] (not shown). These data confirm that Ssb depletion does not lead to a significant stress response, making indirect effects of ssb1/2Δ on [PSI] unlikely.

Overproduced Sup35 protein (8, 9, 19) or Sup35N domain (55) inhibits growth of the [PSI⁺] strains, apparently due to increased accumulation of prion aggregates. Moreover, high-copy-number plasmids, coding for Sup35 or Sup35N, are also toxic for the [psi⁻ PIN⁺] strains, and this toxicity requires the same sequence elements as are required for [PSI] induction, suggesting that toxicity could be related to prion formation (21, 22). We have checked the effect of the ssb1/2Δ on toxicity of overproduced Sup35 and Sup35N. Surprisingly, growth-inhibitory effects of the overproduced Sup35 in [PSI⁺] strains (Fig. ​(Fig.4A),4A), as well as those of the hyperamplified SUP35 and SUP35N plasmids in the [psi⁻ PIN⁺] strains (Fig. ​(Fig.4B),4B), were both decreased rather than increased in an Ssb⁻ background. Thus, Ssb effects on [PSI] formation and toxicity are opposite each other. One could suggest that the decrease in toxicity of the overproduced Sup35 in an Ssb⁻ background contributes to the increase in [PSI] induction that we have observed. However, ssb1/2Δ influenced the growth of moderate Sup35 overproducers, used in [PSI] induction experiments, only slightly (Fig. ​(Fig.2E).2E). Moreover, a 1-order-of-magnitude difference in percentage of [PSI⁺] cells was observed between the Ssb⁺ and Ssb⁻ cultures that had undergone equal numbers of cell divisions (Fig. ​(Fig.2D2D and E). Besides, spontaneous [PSI] formation in the absence of Sup35 overproduction was also greater in the Ssb⁻ strain than in the Ssb⁺ strain (Fig. ​(Fig.2A2A and Table ​Table2),2), while growth rates of the [PSI⁺] strains which did not overproduce the Sup35 protein were not increased by ssb1/2Δ (Fig. ​(Fig.5B).5B). This finding suggests that differences in toxicity of the overproduced Sup35 or Sup35N between Ssb⁺ and Ssb⁻ strains are not sufficient to explain the difference in frequency of [PSI] formation.