Plasmid and strain constructs, PRF, and transcriptional derepression assays.

A YCp50-based S. cerevisiae genomic library was purchased from the American Type Culture Collection (58). The genomic clone that complemented the mof6-1 ts⁻ phenotype was given the name pJD155. Subclones of pJD155 were generated by partial digestion with EcoRI and self ligated by using a Roche rapid ligation kit. The pRS series of plasmids have been previously described (9, 63). Full-length RPD3, PEX6, and AAD14 genes were amplified from genomic DNA from JD932D by PCR using the oligonucleotide primers 1 and 2 (rpd3 alleles), 3 and 4 (PEX6), and 5 and 6 (AAD14) (Table ​(Table2)2) and cloned into pRS314 and pRS316 and were then designated pRPD3, pPEX6, and pAAD14, respectively. The mof6-1 allele was amplified from genomic DNA obtained from strain JD469-2C by using primers 1 and 2. PCRs using the oligonucleotide primers 1 and 2 were carried out under the following conditions: denaturation of dsDNA for 30 s at 95°C, annealing at 48°C for 45 s, and elongation for 4 min. PCR products were purified by using a Qiagen kit, digested with XhoI and PstI, and ligated into pRS314 or pRS316 (63). PCRs using oligonucleotide primers 3 and 4 (PEX6) or 5 and 6 (AAD14) were carried out under the following conditions: denaturation of dsDNA for 30 s at 95°C, annealing for 45 s at 48°C, and elongation for 6 min. PCR products were purified by using a Qiagen kit, digested with KpnI and SalI (PEX6) or XhoI and PstI (AAD14), and ligated into pRS314. To make pHDAC1, the 1.5-kb BamHI fragment containing the HDAC1 cDNA (a generous gift from S. L. Schreiber) was excised from pBJ5/HDAC1-F (70) and inserted into BamHI-digested pG-1 (61), thus placing the human gene under control of the constitutive yeast PGK1 promoter. The synthetic oligonucleotide H151A (Table ​(Table2)2) was used to create pH151A by use of standard methods (43). Subcloning PEX6 into pRS306 (63) generated plasmid pPEX6, which was used to integrate URA3 into the PEX6 locus of yeast strain LNY95. pRPD3, pSIN3, and pSAP30 were generous gifts from M. Hampsey.

Plasmids p−1 and p0, which were used to monitor PRF, have been described previously (73). Briefly, in these plasmids, transcription is driven from the yeast PGK1 promoter into an AUG translational start site. The E. coli lacZ gene serves as the enzymatic reporter, and transcription termination utilizes the yeast PGK1 transcriptional terminator. In the p0 plasmids, lacZ is in the 0 frame with respect to the translational start site, and measurement of β-galactosidase activity generated from cells transformed with these plasmids serves to represent the 0-frame controls. In the p−1 series, lacZ is in the −1 frame with respect to the translational start site and is 3′ of the L-A programmed −1 ribosomal frameshift signal such that β-galactosidase can only be produced as a consequence of a programmed −1 ribosomal frameshift. The efficiency of PRF was calculated by determining the ratio of β-galactosidase activity produced by cells harboring p−1 to the β-galactosidase activity produced by cells harboring p0 and multiplying this ratio by 100. All assays were performed in triplicate.

Plasmids pAV73 and pAV138 were used to monitor transcriptional derepression in cells harboring alleles of RPD3 (76). pAV73 is a URA3 2μm CYC-LacZ fusion reporter plasmid that is used as a control to establish a baseline. PAV138 contains a URS1 site from the HOP1 promoter cloned into the XhoI site of pAV73. It represses lacZ expression in a Ume6p/Sin3p/Rpd3p-dependent manner. The pAV73/pAV138 ratios of β-galactosidase activities were used to calculate the ability of the RPD3 variants to derepress lacZ reporter gene transcription as previously described (76). All assays were performed in triplicate.

In vivo [³⁵S]methionine incorporation.

Labeled methionine incorporation assays were performed as previously described (7). Briefly, isogenic rpd3Δ strains containing wild-type pRPD3 or pmof6-1 were grown in 30 ml of medium lacking methionine and tryptophan at 30°C to an OD₅₉₅ of 1.0. Unlabeled methionine was added to a concentration of 50 μM, and [³⁵S]methionine and [³⁵S]cysteine (Expre³⁵S³⁵S Label; NEN Life Science Products) were added to each culture to final specific activities of 1.1 μCi/ml. Samples were harvested at 0 min and at 15-min intervals for 60 min. Incorporation of the [³⁵S] labels was monitored by cold trichloroacetic acid (TCA) precipitation. For each time point, 1.2-ml aliquots were harvested, from which 0.2 ml was used to determine the OD₅₉₅ of the cultures. One milliliter of ice-cold 20% TCA was added to the remaining 1 ml of each aliquot, incubated on ice for 10 min, heated to 70°C for 20 min, and filtered through prewet Whatman GF/C filters. Filters were sequentially washed with 10 ml of ice-cold 5% TCA and 10 ml of 95% ethanol and dried, and the radioactivity of samples was determined by scintillation counter. All time points were taken in triplicate.

Pulse-chase labeling of rRNA.

Pulse-chase labeling with l-[methyl-³H]methionine was carried out on the isogenic wild-type, mof6-1, rpd3Δ, rpd3-H151A, sin3Δ, and sap30Δ strains as previously described (23, 45). Twenty thousand counts per minute per sample were resolved on a 1.2% formaldehyde-agarose gel. Labeled RNAs were transferred to a zeta probe membrane (Bio-Rad), sprayed with En³Hance (Dupont), and exposed to X-ray film.

Polysome and two-dimensional NEPHGE analyses.

For polysome analyses, cytoplasmic extracts, prepared as described by Baim et al. (2), were fractionated on 7 to 47% sucrose gradients buffered with 50 mM Tris-acetate (pH 7.4), 50 mM NH₄Cl, 12 mM MgCl₂, and 1 mM dithiothreitol (DTT). Gradients were centrifuged in an SW41 rotor at 40,000 rpm for 135 min at 4°C, fractionated, and analyzed by continuous monitoring of A₂₅₄ (55). For nonequilibrium pH gradient gel electrophoresis (NEPHGE) analyses, 40S and 60S ribosomal subunits were separated by ultracentrifugation through a 7 to 27% sucrose gradient in the presence of 500 mM EDTA. Purified ribosomal subunits (≈195 μg/sample) were separated by NEPHGE (pH gradient of 3.5 to 11.5% in the first dimension and 12.5% polyacrylamide separating gel in the second dimension) and were visualized by silver staining by the Kendrick Laboratories (Madison, Wis.).

Preparation of tRNAs and of donor and acceptor fragments.

Yeast tRNAs were charged with [¹⁴C]phenylalanine as previously described (32, 50). Briefly, a 400-μl reaction mix composed of 200 μg of yeast tRNA (Sigma), 25 mM Tris-HCl (pH 7.5), 20 mM MgCl₂, 10 mM ATP, 50 nmol of phenylalanine (313 mCi/mmol; NEN), and 50 μl of aa-tRNA synthetase (9 U/ml; Sigma) was incubated for 25 min at 37°C. After addition of 40 ml of 3 M sodium acetate (pH 5.0), the mixture was extracted twice with an equal volume of water-saturated phenol and once with chloroform. It was then precipitated with 2.5 volumes of ethanol at −20°C for 1 h. After centrifugation for 10 min, the pellet was resuspended in 50 ml of 2 mM potassium acetate (pH 5.0). [¹⁴C]Phe-tRNA^Phe was separated from uncharged tRNAs by using DEAE Sephadex as previously described (57). Acetylation of charged tRNAs was performed as previously described (30). Briefly, [¹⁴C]Phe-tRNA^Phe was resuspended in 200 μl of 0.2 M sodium acetate (pH 5.0), followed by addition of 2.5 μl of acetic anhydride. After a 1-h incubation on ice, another 2.5 μl of acetic anhydride was added, and incubation at 0°C was done for an additional hour. The tRNA was precipitated by addition of 2.5 volumes of ethanol. The [¹⁴C]Phe-tRNA^Phe and acetyl-[¹⁴C]Phe-tRNA^Phe were subsequently digested with 500 U of RNase T1 in 200 μl of 0.3 M sodium acetate (pH 5.0) for 1 h at 37°C, and the reaction mixtures were purified by using DEAE Sephadex as previously described (57). The resulting A site-specific [¹⁴C]Phe-CACCA (acceptor) and P site-specific acetyl-[¹⁴C]Phe-CACCA (donor) fragments were used as substrates for the tRNA fragment binding assays.

Purification of ribosomes and tRNA binding assays.

Salt-washed ribosomes were purified as previously described (48, 50). Briefly, yeast cells were grown in 0.5 liters of YPAD overnight, collected by centrifugation, and washed twice with buffer A (20 mM Tris-HCl [pH 7.5], 10 mM MgCl₂, 1 mM DTT, 0.1 mM EDTA, 0.25 M sucrose). The cell pellet was suspended in 20 ml of buffer A, 30 g of glass beads (0.45-mm diameter) were added, and cells were disrupted by vortexing. The yeast lysate was centrifuged twice for 15 min at 15,000 rpm in a Sorvall S34 rotor, and the supernatant was pelleted at 100,000 × g for 3 h. The pellet was suspended in 6 ml of buffer B (20 mM Tris-HCl [pH 7.5], 10 mM MgCl₂, 1 mM DTT, 0.1 mM EDTA, 0.25 M sucrose, 0.5 M KCl) and placed on a cushion of 3 ml of buffer C (20 mM Tris-HCl [pH 7.5], 10 mM MgCl₂, 1 mM DTT, 0.1 mM EDTA, 1 M sucrose, 0.5 M KCl). After centrifugation at 50,000 rpm for 4 h (SW55 Ti), the pellet was dissolved in 1 ml of buffer A. After a clarifying spin for 1 min in a microcentrifuge, OD₂₆₀ readings were taken (1 A₂₆₀ unit = 19 pmol of ribosomes [1]). The protein content of ribosomes was also estimated by using protein assay reagent (Bio-Rad). Ribosomes were suspended in buffer A at a concentration of 4 pmol/μl and stored at −70°C.

The whole-tRNA and tRNA fragment binding assays were performed by following a modification of previously published protocols (32, 50). Briefly, ribosomes (400 pmol) were incubated with 800 pmol of whole tRNAs or donor or acceptor fragments in 500 μl of a buffer containing 70 mM Tris-acetate (pH 7.2), 40 mM magnesium acetate, 0.4 M potassium acetate, and 50 mM NH₄Cl. Ethanol was added to a final concentration of 30%, and 20-μl aliquots were taken during the time course at 24°C. Samples were diluted to 1 ml with cold buffer (50 mM Tris-HCl [pH 7.2], 0.4 M KCl, 40 mM MgCl₂, 30% ethanol), immediately precipitated onto a Millipore filter, washed with 1 ml of the dilution buffer, and counted in a scintillation counter. The reaction mix without ribosomes was used as the control. All assays were performed in triplicate.

Correlation of growth and frameshifting defects in the rpd3 mutants.

It was observed that the initial appearance and subsequent growth of colonies of rpd3Δ cells transformed with the various plasmid-borne mutant alleles of rpd3 was delayed relative to that of cells containing the wild-type gene. Measurement of growth rates revealed significant quantitative differences. In logarithmic growth, the doubling times of cells harboring either vector alone or the pH151A allele were approximately 1.4-fold longer than that of wild-type controls (doubling time of 3.5 h for cells harboring pRS314 or pH151A versus 2.5 h for cells harboring pRPD3). The growth defect was even greater in mof6-1 cells, where the doubling time was increased approximately 1.6-fold compared with that of isogenic wild-type cells (4.0 versus 2.5 h). Shifting of cells from stationary-phase growth to fresh medium and subsequent monitoring of cell growth rates revealed that the mutants also exhibited significantly different effects on the quality of growth compared with those of isogenic wild-type controls. Particularly striking was the observation that the mutants remained in lag-phase growth for approximately 2 h longer than did wild-type controls (Fig. ​(Fig.3A),3A), suggestive of a defect in the ability of the biosynthetic apparatus to respond to the presence of a rich nutrient source. Similarly, the onset of diauxic shift occurred approximately 2 h earlier for cells harboring pmof6-1 and 1 h earlier for cells harboring pH151A and vector alone than for cells expressing the wild-type gene (Fig. ​(Fig.3A).3A). This result suggests inefficient utilization of carbon source by the mutants.

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FIG. 3.

Correlation of growth and frameshifting defects in the rpd3 mutants. (A) rpd3 mutants display delayed exit from lag phase and early entry into diauxic shift. Stationary-phase cultures of isogenic rpd3Δ strains harboring vector alone (pRS314) or the indicated plasmid-borne alleles of RPD3 were diluted into complete synthetic medium lacking tryptophan to 0.3 U of OD₅₅₀/ml and grown at 30°. At 1-h time points thereafter, aliquots of cells were sampled and OD₅₅₀ readings were determined. Data shown represent the means of each time point through three repetitions of the experiment. Standard deviations were <5%. Solid arrows indicate the approximate point of exit from lag phase, and dotted arrows indicate diauxic shift. (B) Decreased rates of protein synthesis in cells expressing the Mof6-1p form of Rpd3p.[³⁵S]methionine and [³⁵S]cysteine were added to mid-logarithmically growing isogenic rpd3Δ strains containing wild-type pRPD3 or pmof6-1, and samples were harvested at 0 min and at 15-min intervals for 60 min. Incorporation of the [³⁵S] labels was monitored by cold TCA precipitation as described in Materials and Methods. The data were plotted by using the formula y = mx + B, where m is the slope, x is the sample, and B is the y intercept. Rates of protein synthesis correspond to m. All time points were taken in triplicate. (C) Frameshifting defects in the rpd3 mutants are maximized when demand for ribosomes is greatest. Programmed −1 ribosomal frameshifting efficiencies were determined in isogenic rpd3Δ strains harboring vector alone (pRS314) or the indicated plasmid-borne alleles of RPD3 during the three different phases of cell growth. La, lag-phase growth; Lo, log-phase growth; Dx, after diauxic shift. All assays were performed in triplicate. Error bars denote percent error.

Given the original translation-associated defect of mof6-1, we examined whether these cells exhibited any gross defect in protein synthesis. Rates of incorporation of [³⁵S]-labeled methionine and cysteine into newly synthesized protein in mid-logarithmically growing cells were determined as described in Materials and Methods. The results (Fig. ​(Fig.3B)3B) demonstrate that rates of protein synthesis in mof6-1 cells were approximately 75% of that in wild-type cells.

Previous experiments have demonstrated that programmed −1 ribosomal frameshifting efficiencies remain stable throughout the growth cycle in wild-type cells (15). In light of the effects of the mutants on cell growth, −1 PRF efficiencies were monitored during lag phase, log phase, and after diauxic shift in isogenic rpd3Δ cells harboring vector alone, pRPD3, pmof6-1, and pH151A. The results of these experiments show that −1 PRF defects were maximized in the mutants in lag phase, becoming less severe as cells progressed through the growth curve (Fig. ​(Fig.3C).3C). The effect was most notable in mof6-1 cells. The results suggest that the frameshifting defects were maximized in the mutants when demand for newly synthesized ribosomes was the greatest and that −1 frameshifting efficiencies decreased in parallel with the demand for new ribosomes. That no such effect was observed in wild-type cells is in line with the bioeconomic model of regulation of ribosome biosynthesis (80) and suggests a defect in the regulation of ribosome biosynthesis (see below).

Deletion of other genes linked to heterochromatin-associated functions also results in the Mof⁻ phenotype.

Deacetylation of histones by Rpd3p promotes local chromatin condensation, resulting in transcriptional repression of nearby RNA Pol II-transcribed genes (reviewed in reference 10). Although Rpd3p is able to deacetylate histones in vitro, in vivo deacetylation of histones by Rpd3p requires the cofactors. Mutations in any of the components of the Rpd3p/Sin3p/Ume6p repression complex lead to gene-specific derepression of RNA Pol II-regulated genes and concomitant transcriptional activation (33, 40). Conversely, mutations in any of the components of the Rpd3p/Sin3p/Sap30p repression complex lead to the opposite effect, i.e., enhanced transcriptional silencing of RNA Pol II-transcribed genes artificially inserted into heterochromatin contexts (69).

Given the model describing the histone deacetylase complex, if mof6-1 is acting through either of these complexes, then sin3 mutants would also exhibit Mof⁻ phenotypes. Furthermore, if the effect is on a heterochromatin-associated function, e.g., transcription or maturation of rRNAs, then sap30 mutants should also promote increased −1 PRF efficiencies. Conversely, ume6 mutants should promote increased −1 PRF efficiencies if the effect is on genes found in euchromatin, e.g., RP genes. Figure ​Figure44 shows that deletion of either SIN3 or SAP30 resulted in increased −1 PRF efficiencies, while three separate ume6 alleles did not. The sin3Δ and sap30Δ strains also had significant killer virus maintenance defects, whereas ume6 strains were able to stably maintain the killer virus (data not shown). These results demonstrate that (i) the histone deacetylation apparatus is involved in a process that results in a specific translational fidelity defect and (ii) the effect is likely to involve a process in the heterochromatin environment.

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FIG. 4.

Deletion of other genes linked to heterochromatin-associated functions also results in frameshifting defects. The frameshift test vectors p0 and p−1 were introduced into isogenic wild-type (YMH171), rpd3Δ (YMH270), sin3Δ (YMH265), and sap30Δ (YMH277) strains, along with four strains harboring different alleles of UME6 (AJ82, AJ82 11-2, AJ82 66-2, and AJ82 77-2), and programmed −1 ribosomal frameshift efficiencies were determined. All assays were performed in triplicate. Error bars denote percent error.

Mutation of genes involved in the histone deacetylation apparatus results in 60S ribosomal subunit biogenesis defects.

That a defect involving heterochromatin should result in a translational fidelity defect suggested a ribosome biogenesis defect involving rRNA transcription or processing. Given the involvement of the histone deacetylase complex in transcription-associated processes, the effects of these alleles on rRNA transcription and processing were examined by pulsing cells with [³H]methylmethionine, which specifically labels the methylated RNAs, the most abundant of which are those transcribed from the 35S operon. Though no differences were observed with regard to either the rates of 35S pre-rRNA synthesis or its eventual maturation to 18S and 25S rRNAs, the amount of time required for the initial processing step of the 35S pre-rRNA in the mutant cells was delayed by approximately 3 min compared with that for wild-type controls (Fig. ​(Fig.5A).5A). A steady-state analysis revealed that there was no accumulation of any precursors in the mutant cells (data not shown). Polysome analyses of ribosomes isolated from isogenic wild-type, rpd3Δ, mof6-1, rpd3-H151A, sin3Δ, and sap30Δ strains were suggestive of biogenesis defects in the 60S ribosome subunits, as evidenced by decreased levels of 60S ribosomal subunits, increased areas under the 80S peaks, and decreased polysome peaks (Fig. ​(Fig.5B5B).

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FIG. 5.

Mutations of genes involved in histone deacetylation result in ribosome biogenesis defects. (A) Delayed 35S rRNA processing. Pulse-chase labeling with l-[methyl-³H]methionine was carried out on the isogenic wild-type, mof6-1, rpd3Δ, rpd3-H151A, sin3Δ, and sap30Δ strains as previously described (23, 45). Twenty thousand counts per minute per sample were resolved on a 1.2% formaldehyde-agarose gel. Labeled RNAs were transferred to a zeta-probe membrane, sprayed with En³Hance, and exposed to X-ray film. (B) 60S ribosomal subunit biogenesis and polysome defects. Cytoplasmic extracts from isogenic strains were fractionated though sucrose gradients as described in Materials and Methods. Gradients were centrifuged in an SW41 rotor at 40,000 rpm for 135 min at 4°C, fractionated, and analyzed by continuous monitoring of A₂₅₄ (55).

Ribosome biogenesis defects are not due to global defects in RP expression.

It has previously been found that defects in specific RPs result in increased PRF efficiencies (50, 56). Thus, one possible explanation for the observed effects could be that these alleles promote altered expression of RPs. To examine this possibility, approximately 200-μg samples of 60S and 40S ribosomal subunits isolated from isogenic wild-type and mutant cells were separated in two dimensions by NEPHGE, and RPs were visualized by silver staining. No gross differences in the staining patterns were observed between wild-type and mutant samples (data not shown). These results demonstrate that the effects of the rpd3 mutants on PRF and ribosome biogenesis are not due to defects in the synthesis of RPs.

Mutants result in aa-tRNA binding defects.

One previously unexplained phenotype of rpd3 mutants has been their sensitivity to cycloheximide, a translational inhibitor (78). In light of our data showing that these classes of mutants promote a ribosome biogenesis defect specific to 60S subunits, we employed a pharmacogenetic approach using well-characterized antibiotics to investigate the specificity of the defects. Sparsomycin, which increases the affinity of ribosomes for the 3′ (donor) end of peptidyl-tRNAs (34, 39, 51) was used as a P site-specific probe. Anisomycin, which decreases ribosomal affinities for the 3′ (acceptor) ends of aa-tRNAs (6, 29, 62), and paromomycin, which stabilizes binding of near-cognate tRNAs in the decoding center of the small-subunit rRNA (8, 54, 77) served as probes for A site-specific defects. Figure ​Figure6A6A shows that although sparsomycin had no effect relative to the wild type on cells harboring the various rpd3 alleles (rpd3Δ, mof6-1, and rpd3-H151A) or the sin3Δ and sap30Δ mutants, all of the mutants were hypersensitive to anisomycin and all but sap30Δ were hypersensitive to paromomycin. These data indicate that the translational defect caused by mutations in these genes is specific to the ribosomal A site. To further investigate the biochemical basis for these observations, tRNA binding experiments were performed comparing isogenic wild-type and mof6-1 ribosomes. Although no differences were observed in the binding of the 3′ ends of either donor or acceptor fragments (data not shown), the binding profiles for intact aa-tRNAs were significantly different (Fig. ​(Fig.6B).6B). Specifically, mof6-1 ribosomes had decreased initial rates of aa-tRNA binding and had lower overall affinities for aa-tRNAs. In addition, precipitous drop-off in aa-tRNA binding at the 60-min time point suggests that mof6-1 ribosomes are less stable than their wild-type counterparts.

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FIG. 6.

The mutants promote ribosomal A site-specific and peptidyl transfer defects. (A) Pharmacogenetic analyses show hypersensitivity to A site-specific drugs. Tenfold dilutions (from 10⁵ to 10¹ CFU) of isogenic wild-type, rpd3Δ, mof6-1, rpd3-H151A, sin3Δ, and sap30Δ strains were spotted onto selective (H−Trp) medium containing the indicated concentrations of drugs or no-drug controls and incubated at 30°C for 3 days. (B) Ribosomes from mof6-1 cells have decreased binding for aa-tRNA. Ribosomes purified from isogenic wild-type and mof6-1 strains were incubated with molar excess amounts of purified [¹⁴C]Phe-tRNA, and salt-washed ribosomes were diluted at the indicated time points and quickly filtered through nitrocellulose filters. After drying, the filters were counted by liquid scintillation. Assays were performed in triplicate. Data points and error bars indicate means and standard deviations. (C) Ribosomes from mof6-1 cells have decreased peptidyltransferase activities. Time course of the formation of [¹⁴C]phenylalanine-puromycin product in assays using ribosomes isolated from isogenic rpd3Δ cells expressing wild-type or the mof6-1 forms of Rpd3p. Ethyl acetate soluble radioactivity was determined by liquid scintillation counting. Control studies were performed in the absence of puromycin to determine the nonspecific extraction of CACCA[¹⁴C]AcPhe. Control values (generally less than 2%) were subtracted from the values obtained in the presence of puromycin. All experiments were performed in triplicate. Data points and error bars indicate means and standard deviations.

A.M. and J.L.B. contributed equally to this work.

We thank Andrew Vershon, Michael Hampsey, Lenore Neigeborn, and S. L. Shreiber for strains and clones and for helping to provide critical insights.

This work was supported by a grant to J.D.D. from the NIH (GM58859). J.L.B. was supported in part by a training grant from the NIH (T32 AI51967).