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. 2010 May;16(5):1068–1077. doi: 10.1261/rna.2087510

The requirement for the highly conserved G−1 residue of Saccharomyces cerevisiae tRNAHis can be circumvented by overexpression of tRNAHis and its synthetase

Melanie A Preston 1,2, Eric M Phizicky 1,2
PMCID: PMC2856879  PMID: 20360392

Abstract

Nearly all tRNAHis species have an additional 5′ guanine nucleotide (G−1). G−1 is encoded opposite C73 in nearly all prokaryotes and in some archaea, and is added post-transcriptionally by tRNAHis guanylyltransferase (Thg1) opposite A73 in eukaryotes, and opposite C73 in other archaea. These divergent mechanisms of G−1 conservation suggest that G−1 might have an important cellular role, distinct from its role in tRNAHis charging. Thg1 is also highly conserved and is essential in the yeast Saccharomyces cerevisiae. However, the essential roles of Thg1 are unclear since Thg1 also interacts with Orc2 of the origin recognition complex, is implicated in the cell cycle, and catalyzes an unusual template-dependent 3′–5′ (reverse) polymerization in vitro at the 5′ end of activated tRNAs. Here we show that thg1-Δ strains are viable, but only if histidyl-tRNA synthetase and tRNAHis are overproduced, demonstrating that the only essential role of Thg1 is its G−1 addition activity. Since these thg1-Δ strains have severe growth defects if cytoplasmic tRNAHis A73 is overexpressed, and distinct, but milder growth defects, if tRNAHis C73 is overexpressed, these results show that the tRNAHis G−1 residue is important, but not absolutely essential, despite its widespread conservation. We also show that Thg1 catalyzes 3′–5′ polymerization in vivo on tRNAHis C73, but not on tRNAHis A73, demonstrating that the 3′–5′ polymerase activity is pronounced enough to have a biological role, and suggesting that eukaryotes may have evolved to have cytoplasmic tRNAHis with A73, rather than C73, to prevent the possibility of 3′–5′ polymerization.

Keywords: tRNAHis guanylyltransferase, THG1, 3′–5′ polymerization, HTS1, histidyl-tRNA synthetase

INTRODUCTION

Virtually all tRNAHis species have a unique additional guanine nucleotide (G−1) at their 5′ end. This G−1 residue is located at the top of the acceptor stem across from N73, at a position that is normally unoccupied in mature tRNA (Juhling et al. 2009), with the exception of a single tRNAPhe species with a U−1 residue (Schnare et al. 1985). Two distinctly different mechanisms are employed to generate the G−1 residue of tRNAHis. In almost all eubacteria and some archaea (as well as in chloroplasts and most mitochondria), G−1 is encoded in tRNAHis genes opposite C73 and, based on analysis in Escherichia coli, is retained during processing (Orellana et al. 1986). In all sequenced eukaryotes and in other archaea, the G−1 residue is not encoded and is added post-transcriptionally by tRNAHis guanylyltransferase (Cooley et al. 1982), encoded by THG1 in the yeast Saccharomyces cerevisiae (Gu et al. 2003). Thg1 is an essential enzyme that is highly conserved in eukaryotes and in the corresponding archaeal species (Gu et al. 2003). Thg1 adds G−1 opposite A73 in eukaryotes (Cooley et al. 1982; Gu et al. 2003) and opposite C73 in archaeal species (Heinemann et al. 2009; Abad et al. 2010) and in chicken mitochondrial tRNAHis (L'Abbe et al. 1990). The only known tRNAHis species that lack G−1 are found in a small clade of alphaproteobacteria (Wang et al. 2007).

The near universal conservation of the G−1 residue of tRNAHis and the prevalence of two distinctly differing mechanisms for acquiring the G−1 residue suggest that the G−1 residue has crucial roles in the cell. Indeed, one well-documented role of the G−1 residue is for aminoacylation of tRNAHis. The G−1 residue or its phosphate is a strong identity element in vitro for histidylation of E. coli tRNAHis and yeast tRNAHis by their corresponding synthetases (Himeno et al. 1989; Francklyn and Schimmel 1990; Rudinger et al. 1994; Nameki et al. 1995; Fromant et al. 2000; Connolly et al. 2004; Rosen and Musier-Forsyth 2004; Rosen et al. 2006), and in vivo in yeast, based on the concomitant loss of G−1 from tRNAHis and accumulation of deacylated tRNAHis that occurs as Thg1 is depleted (Gu et al. 2005). Thus, the G−1 residue may have been retained during evolution primarily as a determinant for histidyl-tRNAHis synthetases. However, the high conservation of the G−1 residue by two completely different mechanisms is more consistent with the idea that the G−1 residue might itself have an important role in tRNAHis function, and that the synthetases may have evolved afterward to recognize this feature.

Thg1 is also implicated in other important processes. Thg1 may have a role in the cell cycle in yeast, based on the observation that a thg1ts mutant has a defect in the G2/M transition of the cell cycle, and that THG1 interacts genetically and physically with ORC2 (Rice et al. 2005), a component of the origin recognition complex that is crucial for DNA replication (Bell 2002). A role for Thg1 in the cell cycle may also be conserved in other eukaryotes, since the human Thg1 ortholog is cell cycle regulated, and its knockdown leads to defects in cell proliferation and the formation of polycentrosomal cells with multiple spindle poles (Guo et al. 2004). Remarkably, Thg1 also has a distinct 3′–5′ (reverse) polymerase activity in vitro, in which it adds multiple guanosine or cytidine nucleotides in a template-dependent manner to the 5′ ends of different tRNA species that are activated by adenylylation or triphosphorylation (Jackman and Phizicky 2006b). Since either ribonucleotides or deoxynucleotides can be added in the 3′–5′ polymerase reaction, and since the reaction occurs with similar efficiency to the normal G−1 addition that occurs on tRNAHis (Jackman and Phizicky 2006b), this suggests that Thg1 3′–5′ polymerase activity may have a distinct cellular role. However, 3′–5′ polymerase activity has not been observed in vivo.

Here, we address the roles of Thg1 and the G−1 residue of tRNAHis. Remarkably, we find that the normally lethal phenotype of thg1-Δ strains is bypassed by overexpression of both histidyl-tRNA synthetase (HTS1) and tRNAHis. This result demonstrates unequivocally that, despite its widespread conservation, the G−1 residue of tRNAHis is not essential in vivo. Furthermore, this result shows that the only essential Thg1 function is its G−1 addition activity. Surprisingly, thg1-Δ strains overexpressing tRNAHis C73 variants appear healthier than the corresponding strains overexpressing wild-type tRNAHis A73, although cytoplasmic tRNAHis of eukaryotes only has A73. However, we find that tRNAHis C73 is subject to 3′–5′ polymerization in THG1+ strains, providing evidence that 3′–5′ polymerization can occur in vivo, and suggesting a possible explanation for the conservation of A73, rather than C73, in cytoplasmic tRNAHis species.

RESULTS

Development of a sensitive histidine-specific assay to measure tRNAHis function

To study the function of the G−1 residue of tRNAHis in the absence of Thg1 G−1 addition activity, we took advantage of the observation that the anticodon of tRNAHis is necessary and sufficient for tRNAHis guanylyltransferase activity by Thg1 (Jackman and Phizicky 2006a), and devised a tRNAHis nonsense suppressor screen. Since Thg1 specifically recognizes the tRNAHis GUG anticodon, and a tRNAHis nonsense suppressor has a different anticodon (Fig. 1A), this assay should mimic conditions where Thg1 G−1 addition activity is absent, while preserving Thg1 in case its other reported roles are essential.

FIGURE 1.

FIGURE 1.

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Construction of a histidine-specific nonsense suppression system based on Kex2 serine protease. (A) Secondary structure of tRNAHis from S. cerevisiae. tRNAHis mutations described in this study are denoted. RNase A cleavage site used for 5′ end analysis is also indicated. (B) Schematic of Kex2 histidine-specific nonsense suppression assay.

To ensure histidine-specific nonsense suppression, we used Kex2, a serine protease required for yeast mating (Leibowitz and Wickner 1976; Bevan et al. 1998). Since the lack of mature α-factor in kex2 mutants causes a 106-fold reduction in the frequency of mating (Bevan et al. 1998), suppression of a kex2 nonsense mutant provides a convenient and highly sensitive assay to measure nonsense suppression. Moreover, since Kex2 is a serine protease, the required catalytic triad residues (H213, S385, and D175) are each crucial for activity (Fuller et al. 1989), and therefore suppression of a kex2-H213am nonsense mutant requires histidine insertion (Fig. 1B).

We evaluated the Kex2 mating assay by quantifying suppression of kex2 mutants bearing nonsense mutations in the catalytically important S385 and H213 residues, using known suppressors. Control experiments establish that, as expected, mating is undetectable (0.00%–0.01%) in kex2-Δ, kex2-S385am, or kex2-S385oc catalytic triad mutants, and in a control kex2-Y237oc mutant (Table 1). As also expected, suppression of a kex2-S385oc mutant and a kex2-S385am mutant is only observed with the corresponding ochre or amber suppressor tRNA. Consistent with serine-specific suppression, the mating defect of the kex2-S385oc mutant is efficiently suppressed only by the serine-inserting SUQ5-o1 suppressor (2.60%), but not by the tyrosine-inserting ochre suppressor SUP4° (0.00%). Since SUP4° efficiently suppresses the control kex2-Y237oc mutant (18%), the failure of SUP4° to suppress the kex2-S385oc mutant is almost certainly due to insertion of tyrosine, and not serine, at this site. Thus, these results demonstrate that suppression of a kex2-S385oc mutant requires insertion of the catalytic serine residue from the corresponding ochre suppressor. Furthermore, since the SUP4° suppressor is unable to suppress a kex2-H213oc mutant relative to its vector control (0.06% vs. 0.15%), we conclude that suppression of kex2-H213 nonsense mutants provides an effective test for histidine insertion at this catalytically important site.

TABLE 1.

Validation of a histidine-specific nonsense suppression system based on Kex2 serine protease

graphic file with name 1068tbl1.jpg

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Overexpression of the histidyl-tRNA synthetase HTS1 increases the function of tRNAHis am suppressors

Our initial evaluation of tRNAHis am suppressors (Table 1) revealed no measurable suppression of a kex2-H213am mutant when the strains overproduce tRNAHis amber (am) variants containing A73 or C73, the residues normally found at this position in cytoplasmic and mitochondrial tRNAHis, respectively. This result confirms that these tRNAHis am variants are not substrates for the G−1 addition activity of Thg1 in vivo, and suggests that the tRNAHis am variants are not charged with histidine because of the importance of the G−1 residue as an identity element for the histidyl-tRNA synthetase Hts1 (Rudinger et al. 1994; Nameki et al. 1995; Gu et al. 2005; Rosen et al. 2006). In support of this explanation, we find that overproduction of Thg1 increases mating of strains overexpressing tRNAHis am A73, from 0.01% to 5.5% (Table 1). We attempted to increase histidylation of the tRNAHis am suppressors by use of a tRNAHis am G−1–C73 variant containing an encoded G−1 residue (as found in bacteria), but this variant also fails to suppress the mating defect of a kex2-H213am mutant (Table 1), presumably because the G−1 residue is removed during maturation (Orellana et al. 1986; Burkard et al. 1988). However, we find that overexpression of HTS1 dramatically increases mating of kex2-H213am mutant strains expressing tRNAHis am A73 (to 1.3%) or tRNAHis am C73 (to 30.5%), but only marginally increases mating of strains expressing tRNAHis am G73 or tRNAHis am U73 variants (Fig. 2A).

FIGURE 2.

FIGURE 2.

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Overexpression of HTS1 increases suppression by tRNAHis am suppressor variants. (A) Mating assay to measure suppression by tRNAHis am variants. MATα kex2-H213am strains containing tRNAHis am suppressor variants were transformed with 2μ LEU2. HTS1 or a vector control, and mating efficiency was assessed by semi-quantitative and quantitative (right-hand column) mating assays. (B) Nonsense suppression of lys2-801am by tRNAHis am suppressor variants. Strain SS328 (relevant genotype: lys2-801am) was transformed with 2μ URA3 plasmids expressing tRNAHis am variants, 2μ HIS3 HTS1 plasmids, or vector controls, and cells were grown overnight in SD–Ura–His media, serially diluted, and plated on SD–Ura–His media to measure growth after 3 d at 30°C, and on SD–Lys media to measure nonsense suppression after 3 or 8 d.

A control nonsense suppression assay suggests that tRNAHis am suppressors are not misacylated at appreciable levels, since we find that suppression of lys2-801am mutants is undetectable in strains overexpressing tRNAHis am suppressors with A73, G73, or U73, while suppression is barely detectable after 8 d in strains overproducing tRNAHis am C73 suppressors (Fig. 2B). Since the lys2-801am allele is efficiently suppressed by insertion of serine (Brandriss et al. 1976), leucine (Raymond et al. 1985), or tryptophan (Kim and Johnson 1988), the lack of suppression by tRNAHis am suppressor variants indicates lack of charging of these tRNAs by other amino acids. Furthermore, the dramatically increased lys2-801am suppression upon overexpression of HTS1 indicates that histidine is inserted under these conditions. Thus, our results demonstrate that tRNAHis am A73 and C73 variants are each functional for histidine insertion, and suggest that this tRNAHis am function occurs in the absence of the G−1 residue.

The essential function of Thg1 can be suppressed by overexpression of both tRNAHis and HTS1

Because overexpression of HTS1 drastically improves the function of overproduced tRNAHis am variants, we reasoned that overexpression of HTS1 might similarly improve the function of overproduced wild-type tRNAHis(GUG) lacking G−1, and allow a thg1-Δ strain to grow. A thg1-Δ strain will only survive under these conditions if the only essential function of Thg1 is its tRNAHis G−1 addition activity. To address this question, we transformed a thg1-Δ strain containing a URA3 plasmid expressing THG1 under PGAL control (Gu et al. 2003) with plasmids overexpressing tRNAHis(GUG) variants and HTS1, and then streaked transformants on media containing glucose and 5-fluoroorotic acid (5-FOA) to test for growth after loss of the [URA3 PGAL THG1] plasmid (Fig. 3A).

FIGURE 3.

FIGURE 3.

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Thg1 can be bypassed by overexpression of both tRNAHis(GUG) variants and HTS1 in a thg1-Δ strain. (A) Approach for testing the viability of thg1-Δ strains. WG18a (thg1-Δ:kanMX [CEN URA3 PGAL-THG1]) was transformed with a tRNAHis(GUG) variant on a 2μ LEU2 plasmid and a 2μ HIS3 HTS1 plasmid (or vector controls), and the resulting strains were selected on SGal–Leu–His media, and streaked on media containing dextrose and 5-FOA to repress the GAL10 promoter and select against the THG1 covering plasmid. (B) thg1-Δ strains that overexpress tRNAHis variants and HTS1 are viable. Strains were constructed and tested as described in A. (C) thg1-Δ strains have a growth defect. thg1-Δ strains expressing tRNAHis and HTS1 and a wild-type strain were grown, diluted to uniform OD600, and spotted on YPD at the temperatures indicated and grown for 23 h (33°C, 35°C), 41 h (30°C), 67 h (27°C), or 94 h (18°C).

Remarkably, thg1-Δ strains overexpressing HTS1 and either wild-type tRNAHis A73 or tRNAHis C73 are viable after loss of the THG1 gene (Fig. 3B). Viability of the thg1-Δ strains requires the presence of multi-copy plasmids overexpressing both HTS1 and tRNAHis (Fig. 3B). However, we find that thg1-Δ strains are somewhat less healthy than wild-type cells. As measured by growth on plates, thg1-Δ strains overexpressing tRNAHis A73 and HTS1 grow poorly at 30°C and 33°C, and have a severe growth defect at 18°C and 27°C, whereas thg1-Δ strains overexpressing tRNAHis C73 (or tRNAHis G−1–C73) and HTS1 have a distinct but milder growth defect at all temperatures, relative to wild-type cells (Fig. 3C). thg1-Δ strains also exhibit a growth defect in liquid media. At their optimal growth temperature of 33°C in rich media, thg1-Δ strains expressing tRNAHis A73 and tRNAHis C73 have generation times of 2.40 and 1.61 h, respectively, compared to 1.35 h for the wild-type control (Table 2). Another set of thg1-Δ strains expressing HTS1 and tRNAHis C73 or tRNAHis A73 (with a thg1-Δ∷MET15MX rather than a thg1-Δ∷kanMX marker) has a similar set of growth defects, both on plates (data not shown) and in liquid media (Table 2), and the growth defects on plates are complemented by THG1 (data not shown).

TABLE 2.

thg1-Δ strains have a growth defect in liquid media at 33°C

graphic file with name 1068tbl2.jpg

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Because of the widespread conservation of the G−1 residue of tRNAHis, the viability of thg1-Δ strains was unexpected, suggesting that perhaps there was some other mechanism to add a G−1 residue to the tRNA. To determine if the tRNAHis in viable thg1-Δ strains lacks G−1, we purified tRNAHis, and assayed for the presence of G−1 by labeling the 5′ end, followed by treatment with RNase A and thin layer chromatography. This procedure generates the oligonucleotide p*GpGpCp from tRNAHis with a G−1 residue, and p*GpCp if the G−1 residue is missing (see Fig. 1A; Jackman and Phizicky 2006b). As shown in Figure 4, virtually all of the tRNAHis from wild-type cells contains the G−1 residue (96%), whereas, tRNAHis from thg1-Δ strains lacks any clearly detectable amounts of G−1 residue (<3%). Thus, we conclude that the G−1 residue of tRNAHis is itself not absolutely necessary for life in yeast, as long as the strain has compensating increased levels of both tRNAHis and histidyl-tRNA synthetase.

FIGURE 4.

FIGURE 4.

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tRNAHis species isolated from thg1-Δ strains lack G−1. tRNAHis was purified from thg1-Δ strains or a wild-type strain, 5′ end-labeled with [γ-32P]ATP, and digested with RNase A. 4000 cpm of digested tRNAHis was spotted on a silica thin-layer chromatography (TLC) plate, and resolved in solvent containing n-propanol:NH4OH:H2O (55:35:10) for about 16 h, dried, and radioactivity was visualized using a Storm PhosphorImager. Note: the migration of the solvent front was not uniform in this TLC experiment.

The growth defect of thg1-Δ strains suggests that they may be limited for histidyl-tRNAHis, despite the overproduction of HTS1. We therefore analyzed aminoacylation by Northern analysis of RNA prepared under acidic conditions (Chernyakov et al. 2008). As previously observed in wild-type cells (Gu et al. 2005), tRNAArg is nearly 100% aminoacylated and tRNAHis is only ∼60% aminoacylated (Fig. 5). We find that a thg1-Δ strain overexpressing HTS1 and tRNAHis A73 has somewhat less aminoacylated tRNAHis than wild-type cells, although the precise amount of aminoacylated tRNAHis is difficult to accurately quantify. However, we find that thg1-Δ strains overexpressing tRNAHis C73 (or tRNAHis G−1–C73) have substantially more aminoacylated tRNAHis than wild-type cells (five- and threefold, respectively). The greater than normal levels of aminoacylation in thg1-Δ strains with either tRNAHis C73 or tRNAHis G−1–C73 suggest that some other defect in these strains causes them to grow poorly relative to a wild-type strain.

FIGURE 5.

FIGURE 5.

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tRNAHis variants from thg1-Δ strains have decreased aminoacylation. RNA from wild-type and thg1-Δ strains was isolated in acidic conditions to preserve aminoacylation, and 2 μg of RNA was resolved on an acidic gel, analyzed by Northern blotting for tRNAHis, tRNAArg(ICG), and 5S rRNA as described in Materials and Methods, and quantified using ImageQuant software. A control sample was treated with base (+) to detect migration of deacylated tRNA species in the acidic PAGE gel system. Solid arrows, aminoacyl-tRNA; dotted arrows, uncharged tRNA.

It is surprising that thg1-Δ strains overexpressing both tRNAHis and HTS1 have relatively low ratios of charged to uncharged tRNAHis (5% for tRNAHis A73 and 29%–36% for tRNAHis C73), despite the overproduction of HTS1, since a control experiment shows that the chromosomal HTS1 gene is sufficient for a nearly wild-type ratio of charged to uncharged tRNAHis in strains overexpressing tRNAHis (data not shown). Therefore, it seemed plausible that these low charging ratios might arise from instability of the histidyl-tRNAHis, rather than the lack of sufficient charging by Hts1. Indeed, we previously showed that the ester bond of His-tRNAHis is unusually unstable at pH 4.5, compared to other tRNAs (Chernyakov et al. 2008). To determine if the absence of the G−1 residue exacerbates the instability of His-tRNAHis, we end-labeled purified tRNAHis, charged the tRNA with histidine in vitro, and measured the stability of the histidyl-tRNAHis by monitoring its mobility on acid-urea gels. We find that His-tRNAHis lacking the G−1 residue is nearly as stable as His-tRNAHis containing G−1 in buffer at pH 7.5, and is slightly more stable at pH 4.5 (Fig. 6). We thus presume that absence of the G−1 residue of tRNAHis does not significantly enhance the instability of histidyl-tRNAHis in the cell.

FIGURE 6.

FIGURE 6.

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Lack of G−1 on tRNAHis has no effect on ester bond stability of histidyl-tRNAHis. (A) Histidyl-tRNAHis ester bond stability in pH 7.5 buffer at 37°C. tRNAHis species purified from wild-type or thg1-Δ strains were 5′ end-labeled with 32P and aminoacylated in vitro. Then, His-tRNAHis with or without G−1 was incubated in pH 7.5 buffer at 37°C to assess the aminoacyl-tRNA stability (ester bond stability). The resulting product was run on a denaturing polyacrylamide gel, dried and exposed to a phosphor screen. Percent aminoacylation was calculated as described in Figure 5. Solid arrow, histidyl-tRNAHis; dotted arrow, uncharged tRNAHis. (B) Histidyl-tRNAHis ester bond stability in pH 4.5 buffer at 37°C. Histidyl-tRNAHis species were treated essentially as described in A, except that each was incubated in pH 4.5 buffer for the specified times prior to acidic PAGE analysis.

Thg1 3′–5′ reverse polymerization of tRNAHis C73 occurs in vivo

The more efficient growth of thg1-Δ strains expressing tRNAHis C73 instead of tRNAHis A73 suggests that tRNAHis C73 is a more efficient substrate than tRNAHis A73 for Hts1 in vivo, which is consistent with earlier in vitro analysis of tRNAHis determinants of yeast Hts1 (Nameki et al. 1995). This preference for tRNAHis C73 raises the question of why cytoplasmic tRNAHis species have A73 rather than C73. Since tRNAHis C73 is a substrate for the 3′–5′ polymerase activity of Thg1 in vitro (Jackman and Phizicky 2006b), we reasoned that tRNAHis C73 might also be a substrate for 3′–5′ polymerization in vivo. As shown in Figure 7, tRNAHis in strains overexpressing tRNAHis C73 and THG1 has substantial levels of G−2-containing ends (23.4%), and wild-type strains overexpressing tRNAHis C73 have distinct levels of G−2-containing ends (2.4%). This result demonstrates unequivocally that tRNAHis C73 is subject to 3′–5′ polymerization by Thg1 in vivo. Moreover, the actual efficiency of 3′–5′ polymerization of tRNAHis C73 in vivo is underestimated because the purified tRNAHis C73 is unavoidably contaminated with endogenous tRNAHis A73 (which does not have detectable levels of G−2) due to lack of discrimination by the biotinylated oligomer used for tRNAHis purification.

FIGURE 7.

FIGURE 7.

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Thg1 catalyzes reverse polymerization of tRNAHis C73 in vivo. tRNAHis was purified from wild-type strains overexpressing THG1 and a tRNAHis variant or the corresponding vector controls, as indicated, and the 5′ end of the tRNAHis was labeled and analyzed after RNase A digestion as described in Figure 4. Standards (pGGGC, pGGC, and pGC) were made by labeling the 5′ end of synthetic oligomers containing the corresponding sequence at the 5′ end, followed by RNase A digestion, and control tRNAHis transcripts that contained G−1 (76-mer) or lacked G−1 (75-mer), were treated the same way. The % G−2 was calculated from the quantification of the PhosphorImager values, using ImageQuant.

DISCUSSION

We have provided two lines of evidence demonstrating that tRNAHis lacking its G−1 residue can function in yeast. First, we showed that tRNAHis am nonsense suppressors can suppress a kex2-H213am mutant. The tRNAHis am suppressors almost certainly function in the absence of G−1 since mutation of the GUG anticodon drastically reduces Thg1 G−1 addition activity in vitro (Jackman and Phizicky 2006a). Moreover, suppression almost certainly occurs by insertion of histidine, since the Kex2 serine protease requires histidine at the catalytic H213 position, and since suppression requires overexpression of HTS1. Furthermore, it is unlikely that either tRNAHis am C73 or tRNAHis am A73 is appreciably mischarged, since suppression of a lys2-801am mutation is almost undetectable, although this mutation is known to be suppressed by insertion of any of several different amino acids (Brandriss et al. 1976; Raymond et al. 1985; Kim and Johnson 1988), and since almost all the observed suppression requires overexpression of HTS1. Second, we showed that thg1-Δ cells are viable in spite of the lack of detectable G−1 in their tRNAHis, but only if both tRNAHis and HTS1 are overproduced. Thus, despite the near universal conservation of the G−1 residue of tRNAHis in different organisms, the G−1 residue is not absolutely essential for tRNAHis function, at least in yeast. This result is especially surprising in view of the two completely different mechanisms by which the G−1 residue is conserved throughout evolution: by Thg1 G−1 addition activity (Gu et al. 2003; Abad et al. 2010), or by retention of an encoded G−1 residue due to aberrant RNase P processing (Orellana et al. 1986; Burkard et al. 1988).

These results may suggest that the persistence of the G−1 residue of tRNAHis in different organisms is a consequence of an ancient evolutionary trap, in which two different recognition mechanisms evolved for charging tRNAHis. The predominant mechanism requires the G−1 residue or its phosphate as a crucial identity element for the corresponding histidyl-tRNA synthetase (HisRS), based on experiments in E. coli and yeast (Himeno et al. 1989; Francklyn and Schimmel 1990; Rudinger et al. 1994; Nameki et al. 1995; Fromant et al. 2000; Connolly et al. 2004; Rosen and Musier-Forsyth 2004; Gu et al. 2005; Rosen et al. 2006). Since the G−1 residue is unique to tRNAHis, there would be strong evolutionary pressure to maintain this recognition element once it had evolved. The other mechanism occurs in the clade of alphaproteobacteria in which the G−1 residue is not found, and in which the HisRS requires other recognition elements for tRNA charging (Ardell and Andersson 2006; Wang et al. 2007). However, it remains possible that the G−1 residue has another role in the cell, and that recognition of this G−1 residue by HisRS evolved subsequently. For example, the presence or absence of the G−1 residue might be used to modulate tRNAHis function as part of a regulatory response. Alternatively, the G−1 residue might play a role during the accommodation step of translation, based on the distortions that occur in the upper portion of the tRNA acceptor stem during this step (Schmeing et al. 2009), and the known difference in stability of the acceptor stem in the absence of G−1 (Seetharaman et al. 2003). This putative other role of the G−1 residue might be reflected in the growth defect of thg1-Δ strains, or might be revealed by more sophisticated assays of tRNA function in these strains.

Because thg1-Δ strains are viable only if both tRNAHis and HTS1 are overproduced, our results virtually prove that the only essential role of Thg1 is to increase tRNAHis function, presumably by its G−1 addition activity, or conceivably by some other Thg1 activity that enhances tRNAHis function independent of G−1 addition. An essential G−1 addition activity of Thg1 is also consistent with biochemical evidence that the G−1 residue of tRNAHis is a strong determinant for histidylation by Hts1 (Rudinger et al. 1994; Nameki et al. 1995; Gu et al. 2005; Rosen et al. 2006), and with the coincident deacylation of tRNAHis and growth arrest that occurs during Thg1 depletion in vivo (Gu et al. 2003, 2005).

We therefore infer that the other activities attributed to Thg1 are not essential, including its 3′–5′ polymerase activity (Jackman and Phizicky 2006b), its genetic and physical interaction with Orc2 of the origin recognition complex (Rice et al. 2005), and its role in the G2/M phase of the cell cycle (Rice et al. 2005), unless one of these roles is also essential because of a requirement for tRNAHis function. Lack of sufficient functional tRNAHis might, for example, prevent translation of histidine codons of particular essential genes, much as reduced tRNAArg modification in yeast trm9 mutants results in methyl methanesulfonate (MMS) sensitivity and lower expression of specific DNA repair genes (Begley et al. 2007). Alternatively, functional tRNAHis may be essential in some other capacity, much as tRNAs have been implicated in retroviral DNA replication (Jiang et al. 1993). However, since thg1-Δ strains expressing tRNAHis C73 grow substantially slower than wild-type cells despite their abundance of charged tRNAHis, we infer either that these cells have a translation defect due to the alterations of tRNAHis (lack of the G−1 residue or presence of C73), or that Thg1 has another role that is important but not essential for normal growth.

The substantial 3′–5′ polymerization observed in strains overexpressing both tRNAHis C73 and Thg1 and the distinct, but lower, levels of 3′–5′ polymerization observed in strains with endogenous Thg1 emphasize the possibility of a biological function for the 3′–5′ polymerase activity of Thg1, since this activity is so readily detected in vivo. Moreover, it appears that tRNAHis C73 is at least as good a substrate for 3′–5′ polymerization as it is for G−1 addition, based on the disappearance of G+1-containing tRNAHis upon THG1 overexpression, and the concomitant appearance of G−2-containing tRNAHis. This is the first documented case in which Thg1 (or any known gene product) has been shown to catalyze 3′–5′ polymerization in vivo, although this activity is catalyzed in the mitochondria of Acanthamoeba castellani and related organisms by unknown gene products (Lonergan and Gray 1993; Laforest et al. 1997; Price and Gray 1999). We and others have speculated previously that the 3′–5′ polymerase activity of Thg1 might have a role in tRNA repair, or in DNA repair or replication, to account for the reported cell cycle phenotype of a thg1 mutant, or the genetic and physical interactions of Thg1 with Orc2 (Jackman and Phizicky 2006b; Abad et al. 2010), which has been implicated in transcriptional silencing, mitotic spindle assembly, and sister chromatid cohesion (Bell et al. 1993; Foss et al. 1993; Prasanth et al. 2004; Suter et al. 2004; Shimada and Gasser 2007), as well as in DNA replication (Bell 2002). The identification of such roles awaits future experiments.

Our data also provide insight regarding the conservation of A73 rather than C73 in the cytoplasmic tRNAHis of eukaryotes. There is little doubt that C73 is a stronger in vivo determinant for histidylation by Hts1 than is A73, since both aminoacylation and growth of a thg1-Δ strain are more efficient in cells expressing tRNAHis C73 than in those expressing tRNAHis A73, and since tRNAHis am C73 suppressors are more efficient than tRNAHis am A73 suppressors. The increased efficiency of C73 versus A73 as a determinant for histidylation is consistent with previous in vitro analysis of Hts1 from yeast (Nameki et al. 1995). In light of the preference of Hts1 for C73 of tRNAHis, the evolutionary conservation of A73 in the cytoplasmic tRNAHis of eukaryotes might be viewed as somewhat of a mystery, particularly since in archaea and bacteria, C73 is exclusively found in tRNAHis species (Marck and Grosjean 2002). However, the observed 3′–5′ polymerization of tRNAHis C73 by Thg1 suggests that eukaryotes have evolved to have tRNAHis with A73 instead of C73 to protect against the possibility of 3′–5′ polymerization, which would form an additional G−2:C74 base pair, thus perturbing the universally conserved CCA end of tRNA. Based on results in bacteria, this additional tRNA base pair might interfere with a conformational change important for translation (Schmeing et al. 2005).

MATERIALS AND METHODS

Construction of yeast strains

Strains and oligomers used to construct strains are listed in Supplemental Tables S1 and S2. To construct MBY108D (relevant genotype: MATα kex2-Δ), kex2-Δ∷kanMX DNA and flanking sequences were amplified from the corresponding knockout strain (Giaever et al. 2002) (Open Biosystems), and DNA was transformed in BY4742 (MATα), followed by selection on YPD media (Sherman 1991) containing 300 μg/mL geneticin (G418, Gibco).

MBY302A (relevant genotype: MATα The kex2-H213am) was constructed by replacement of the KEX2 locus with a cassette containing the 5′ flanking region of KEX2, followed by URA3, a repeat of the 5′ flanking region, and the kex2 mutated gene. Next, URA3 and the repeated 5′ flanking region were removed by selection on FOA. The cassette was constructed in two steps. First, the URA3 gene was PCR-amplified with oligomers URA3(KpnI) forward and URA3(BamHI) reverse, followed by digestion and ligation into MAB902, which contains the kex2-His213am gene and the 5′ flanking region from −485 (see Supplemental Tables S3, S4). Second, the 5′ flanking region of KEX2 (from −485 to −221) was cloned by PCR amplification with Kex2 −485 Up(SacI) forward and Kex2 −221 Up(KpnI) reverse, followed by restriction digestion and ligation into the plasmid from step 1. The resulting plasmid, MAB868, was digested to release the cassette for transformation into wild-type MATα (BY4742), followed by selection on SD–Ura media. Finally, URA3 and one of the repeated 5′ flanking regions of KEX2 were removed by homologous recombination, by selection on media containing 5-FOA, to generate MBY302A.

thg1-Δ∷MET15MX strains were generated by construction of a MET15MX cassette containing MET15 flanked by ∼250 base pairs of kanMX upstream and downstream DNA (plasmid MAB772), followed by transformation of DNA digested with BamHI and KpnI into thg1-Δ∷kanMX strains.

Construction of plasmids

Oligomers used to construct plasmids are listed in Supplemental Table S3. The 2μ HIS3 vector used in this study (MAB850) was constructed by PCR amplifying the HIS3 gene to insert NcoI and XhoI restriction sites at the ends, followed by ligation into YEplac181 (Gietz and Sugino 1988) that was PCR-amplified to exclude the LEU2 gene and to insert the corresponding restriction sites.

Plasmids containing KEX2, HTS1, and the tRNAHis gene tH(GUG)G2 (and other tRNA genes) were constructed by standard cloning methods after PCR amplification of the genes and flanking regions from appropriate DNA.

Site-directed mutagenesis of Kex2 and tRNA variants

Site-directed mutagenesis was performed using the Quikchange mutagenesis kit (Stratagene), using appropriate primers to generate the corresponding plasmids (Supplemental Table S4).

Isolation of bulk low molecular weight RNA

Wild-type and thg1-Δ strains were grown in YPD at 33°C to an OD600 of 1.5, and harvested in 300 OD pellets, and low molecular weight RNA was isolated using hot phenol, as described previously (Jackman et al. 2003).

tRNA purification

tRNAHis was purified from bulk low molecular weight yeast RNA using a 5′-biotinylated oligomer, 5′ Bio tRNAHis (5′ Bio-GCCATCTCCTAGAATCGAACCAGGG-3′, Integrated DNA Technologies) that is complementary to nucleotides 48–72 of tRNAHis, essentially as described (Jackman et al. 2003), followed by desalting and concentration using Amicon Ultra-4 10,000 MWCO columns (Millipore).

Analysis of aminoacylated RNA

RNA was isolated from 40 OD of cells in acidic conditions (pH 4.5), and 5-μg samples were resolved by acidic PAGE, as described (Chernyakov et al. 2008), followed by transfer to Hybond N+ membrane (Amersham Biosciences), UV cross-linking, and hybridization with 5′-labeled oligomers tRNAHis P1 (5′-GCCATCTCCTAGAATCGAACCAG-3′), ArgP1 (5′-TAGCCAGACGCCGTGAC-3′), and 5S RNA (5′-GGTAGATATGGCCGCAACC-3′) to detect tRNAHis, tRNAArg (ICG), and 5S rRNA, and visualization with a Storm PhosphorImager (Molecular Dynamics).

Quantitative and semi-quantitative mating assays

MATα strains to be tested were grown to OD 0.8 in SD–Ura–Leu media, resuspended in YPD to 1 OD/mL, serially diluted 10° to 104-fold, and mixed with an equal volume of BY4741 MATa cells at 1 OD/mL, and incubated at 30°C for 3.5 h. Then cells were either spotted (semi-quantitative assay) or spread (quantitative assay) onto SD–Met–Lys–Leu–Ura plates to select both for mating and for plasmids expressed in the MATα strains. Mating efficiency was normalized to the mating efficiency of wild-type MATα cells, which varied from ∼30% to 100%.

5′ end labeling of tRNA

Purified tRNAHis (20 pmol) was treated with calf intestinal phosphatase (Roche) to remove the 5′ phosphate, phenol-extracted, and ethanol-precipitated, and resuspended tRNA was labeled with 25 pmol of [γ-32P]ATP (7000 Ci/mmol MP Biologicals) using T4 polynucleotide kinase (Roche), followed by removal of excess label using a Micro Bio-spin 6 chromatography column (Bio-Rad), and gel purification.

In vitro aminoacylation

His6-Hts1 was purified by immobilized metal ion affinity chromatography (Jackman et al. 2003), and 5′ end-labeled tRNAHis species from wild-type or thg1-Δ [2μ LEU2 tRNAHis A73, 2μ HIS3 HTS1] strains were incubated in a buffer containing 50 mM HEPES (pH 7.5), 4 mM DTT, 20 mM KCl, 10 mM MgCl2, 2.5 mM ATP, 1 mM L-Histidine, and 2.5 μM yeast HisRS (His6-Hts1) for 30 min at 30°C, essentially as described (Rosen et al. 2006). Then, samples were treated with phenol saturated with 0.3 M NaOAc (pH 4.5), ethanol-precipitated, and resuspended in cold 10 mM NaOAc (pH 4.5), 1 mM EDTA. Aminoacylated tRNAHis was stored at −20°C.

Aminoacyl-tRNA stability assay

In vitro aminoacylated, 5′ end-labeled tRNAHis was incubated in pH 7.5 buffer (50 mM PIPES [pH 7.5], 20 mM KCl, 10 mM MgCl2) or pH 4.5 buffer (50 mM PIPES [pH 4.5], 20 mM KCl, 10 mM MgCl2) at 37°C for various times to assess ester bond stability, and tRNA was ethanol-precipitated, resuspended in 10 mM sodium acetate (pH 4.5), 1 mM EDTA, and resolved by acidic PAGE, followed by gel drying and quantification.

SUPPLEMENTAL MATERIAL

Supplemental material can be found at http://www.rnajournal.org.

ACKNOWLEDGMENTS

We thank Elizabeth Grayhack for invaluable insight and discussions throughout this work. We also thank Jeffrey Zuber and Michael Guy for technical assistance during preparation of the figures. This research is supported by NIH Grant GM52347 to E.M.P. M.A.P. was supported by NIH Training Grant in Cellular, Biochemical and Molecular Sciences 5T32 GM068411.

Footnotes

Article published online ahead of print. Article and publication date are at http://www.rnajournal.org/cgi/doi/10.1261/rna.2087510.

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