|Home | About | Journals | Submit | Contact Us | Français|
We report that Esf1p (Ydr365cp), an essential, evolutionarily conserved nucleolar protein, is required for the biogenesis of 18S rRNA in Saccharomyces cerevisiae. Depletion of Esf1p resulted in delayed processing of 35S precursor and a striking loss of 18S rRNA. Esf1p physically associated with ribosomal proteins and proteins involved in 18S rRNA biogenesis. Consistent with its role in 18S rRNA biogenesis, Esf1p also physically associated with U3 and U14 snoRNAs, but did not appear to be a core component of the SSU processome. These data indicate that Esf1p plays a direct role in early pre-rRNA processing.
Ribosome biosynthesis is one of the major metabolic activities in cells, and is highly conserved from yeast to humans. It occurs mainly in the nucleolus, a specialized compartment in the nucleus. Current knowledge of ribosome biogenesis is largely derived from studies of the yeast Saccharomyces cerevisiae (1,2). This process starts with the transcription of ribosomal RNA (rRNA) genes by RNA polymerase I and III into 35S pre-rRNA and 5S pre-rRNA. The 35S precursor is extensively modified and cleaved into the mature 18S, 5.8S and 25S rRNA through the coordinated action of a variety of endonucleases, exonucleases, RNA helicases and other protein factors (Fig. (Fig.11).
The U3 small nucleolar RNA (snoRNA), together with associated proteins, plays a central role in the 18S rRNA biogenesis (1,2) and is essential for the early cleavage events at sites A0, A1 and A2 (1–3). Recently, protein complexes that contain the U3 snoRNA and participate in the biogenesis of 40S ribosome subunits have been identified. Dragon and colleagues identified a large nucleolar ribonucleoprotein (RNP), called the small subunit (SSU) processome, required for 18S ribosomal RNA biogenesis (4). It contains the U3 snoRNA and at least 28 proteins, including 17 polypeptides (Utp1–17, for ‘U three protein’) that had not previously been shown to be associated with U3 or implicated in pre-rRNA processing events. We have recently discovered that the SSU processome consists of at least three distinct ‘sub-complexes’ and each physically associates with U3 snoRNA (5). Grandi and colleagues reported 90S pre-ribosomes formed at very early stages of ribosome biogenesis (6). The purified 90S pre-ribosomes contained not only the U3-specific proteins, but also other nucleolar proteins with a known role in 18S rRNA processing and 40S subunit assembly and additional uncharacterized proteins. However, seven components identified in the SSU processome (Snu13p, Utp3p, Utp5p, Utp7p, Utp11p, Utp14p and Utp16p) were not found in the isolated 90S pre-ribosomes.
In addition to the U3 snoRNA, snoRNAs U14, snR10 and snR30 are required for early cleavages at sites A1 and A2 and hence also contribute to 18S rRNA biogenesis (7–9). However, the protein complexes containing these RNAs are less well-characterized than the U3-containing SSU processome. Furthermore, despite the application of several proteomic approaches to characterize early pre-ribosomal protein complexes (4–6) it is likely that not all proteins involved have been detected (4).
We recently carried out a large-scale survey of non-coding RNA processing phenotypes in yeast mutants using a microarray-based approach (10). A mutated version of YDR365c, a gene that is co-regulated with established rRNA processing factors at the transcriptional level (11), displayed an array phenotype suggesting involvement in ribosome biogenesis (10). Ydr365cp has not been identified in the protein complexes described above, although it has been associated with different rRNA processing factors in other large-scale studies (12,13). Here, we provide genetic and biochemical evidence that YDR365c is required for the early cleavages at sites A0, A1 and A2 that lead to 18S rRNA synthesis, although it does not appear to be a core component of the SSU proces some. Owing to its apparently direct role in 18S rRNA biogenesis, we have named the gene ESF1 (Eighteen S rRNA Factor 1).
The S.cerevisiae strains used for phenotypic analysis in this study are R1158 (wild type) (14) and TH_2070, the tetO7-ESF1 mutant (10). The vector pBS1479 (15) was used to C-terminally tag Esf1p in strain YCK245 (a derivative of W303-1A) (16), creating strain YNJK654. pRS411 (a plasmid containing MET15) was a gift from Jef Boeke. All yeast cultures were at 30°C. Yeast transformation was carried out according to a protocol described previously (17). Rich medium YPD (1% yeast extract, 2% peptone, 2% dextrose) (18) was used to culture yeast for protein purification. Synthetic medium SD (18) was used to culture yeast for RNA isolation and pulse–chase labeling.
For depletion of Esf1p protein, tetO7-ESF1 (and the isogenic wild-type control strain) was exposed to 10 µg/ml doxycycline (Sigma) for a total of 24 h before harvesting for RNA extraction. RNA extraction and northern blotting were performed as described previously (10). Oligonucleotides specific for 35S pre-rRNA are: 18S, 5′-CAGAAGGAA AGGCCCCGTTGGAAATCCAGTACACGAAA AAATCG GACCGG-3′; 25S, 5′-TTCCCAAACAACTCGACTCTTCG AAGGCACTTTACAAAGAACCGCACTCC-3′; DA2, 5′-GAAAGAAACTTACAAGCCTAGCAAGACCGCGCACTTAAGCGCAGGCCCGG-3′; A2A3, 5′-TACCTCTGGG CCCCGATTGCTCGAATGCCCAAAGAAAAAGTTGCAAAGAT-3′; EC2, 5′-TCCAATGAAAAGGCCAGCAATT TCAAGTTAACTCCAAAGAGTATCACTCAC-3′; 5′-ETS-A0, 5′-GGAAATGCTCTCTGTTCAAAAAGCTTTTACA CTCTTGACCAGCGCACTCC-3′. Oligonucleotides for snoRNAs are: U3, 5′-CCCTATCCCTTCAAAAAAGAA GTACATAGGATGGGTCAAGATCATCGCGC-3′; U14, 5′-GCGGTCACCGAGAGTACTAACGA-3′; snR10, 5′-CA CATTCTTCATGGGTCAAGAACGCCCCGG-3′; snR30, 5′-TCCATATATATCATGGCAACAGCCCCCGAA-3′.
R1158 (wild type) and tetO7-ESF1 mutant were transformed with pRS411 and grown to 0.8 × 107 cells/ml over a period of 24 h in SD-Met medium containing 10 µg/ml doxycycline. Pulse–chase labeling was performed as described by Kressler et al. (19) with minor modifications. Aliquots comprising of 20 000 c.p.m. were loaded on each lane of a 1% agarose/glyoxal gel and RNA was transferred to a Hybond-N+ nylon membrane (Amersham Pharmacia) by overnight downward capillary transfer. Membranes were sprayed with EN3HANCE (PerkinElmer Life Sciences), dried, and exposed to X-ray films at –80°C with an intensifying screen and visualized by autoradiography.
Four liters of YNJK654 cells were grown in YPD to 1.5 × 107 cells/ml, then harvested. The cell pellets were washed twice with cold water and frozen with liquid nitrogen. Frozen cell pellets were broken with dry ice in a small coffee grinder (Krups), and 10 ml IPP150 buffer (10 mM Tris pH 8.0, 150 mM NaCl and 0.1% Triton X-100) plus 1 mM DTT and protease inhibitors were added to the lysed cells. The broken cells were subjected to centrifugation at 13 000 r.p.m. in an SS-34 rotor (20 200 g) for 1 h at 4°C. The lysates were mixed with 200 µl IgG agarose beads for 2 h at 4°C. After five washes with 0.5 ml IPP buffer, the IgG beads were collected and the RNA associated with the beads extracted with hot phenol and RNase-free DNase I was used to remove DNA in the samples as described (10). For the purification of protein complexes, the eluates from IgG columns were further purified with calmodulin as previously described (16). The purified proteins were separated by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) on gels containing 10% polyacrylamide, visualized by silver staining and identified by MALDI-TOF mass spectrometry.
ESF1 encodes a protein of 628 amino acids with a calculated molecular weight of 72.4 kDa and pI of 4.99. The encoded Esf1 polypeptide has a coiled-coil region located between residue 426 and 492, and a bipartite nuclear localization signal near its N-terminus (from residue 447 to 464). We identified clear sequence counterparts in other organisms, including Schizosaccharomyces pombe, Arabidopsis thaliana, Drosophila melanogaster, Neurospora crassa, Drosophila melanogaster, Mus musculus and Homo sapiens. The multiple sequence alignment reveals what appear to be several discrete conserved regions (Fig. (Fig.2).2). In a systematic deletion analysis, Giaever and colleagues (20) reported that ESF1 is essential for viability. The protein has been localized to the nucleus, possibly enriched in the nucleolus, by three independent studies (21–23).
Our previous microarray study indicated that the tetO7-ESF1 mutant had a defect in 18S rRNA biogenesis (10). To investigate this further, we analyzed the steady-state levels of mature and precursor rRNA molecules by northern blotting. Probes specific for mature 18S and 25S rRNAs confirmed that the level of 18S rRNA was dramatically reduced upon depletion of Esf1p, relative to 25S rRNA (Fig. (Fig.3).3). Hybridizing the membrane with oligonucleotides complementary to D-A2 and A2-A3 regions (probes DA2 and A2A3) revealed that 20S pre-rRNA (the direct precursor to 18S rRNA) and the 27SA2 pre-rRNA (normally a precursor to the 25S rRNA), were not detected in the tetO7-ESF1 mutant. We also observed accumulation of 35S pre-rRNA and appearance of aberrant 23S pre-rRNA in the mutant. The 23S pre-rRNA is generated by cleavage at A3 in the 35S precursor without the early cleavages at sites A0, A1 and A2 (2). These data indicate that ESF1 is required for pre-rRNA processing at sites A0, A1 and A2 that lead to production of the mature 18S rRNA.
To further confirm that depletion of Esf1p affects pre-rRNA processing, we carried out pulse–chase labeling of the pre-rRNA with [methyl-3H]methionine. In wild-type cells, the 35S precursor was processed into 25S and 18S rRNAs in ~5 min (Fig. (Fig.4).4). In contrast, tetO7-ESF1 mutant contained very little 20S pre-rRNA and 18S rRNA at any time point examined (up to 15 min). The formation of 25S rRNA was delayed, but it was produced much more rapidly than 18S rRNA. Processing of the 35S precursor was also delayed in the Esf1p-depleted strain, perhaps because cleavage at A3 in the absence of cleavage at A2 is not the primary pathway. This would also explain the delay in 25S formation (Fig. (Fig.4),4), and the slight reduction in steady-state 27S levels (Fig. (Fig.33).
To assess whether Esf1 is involved in the biogenesis or maintenance of the snoRNAs U3, U14, snR10 and snR30 [required for cleavage at A0, A1 or A2 (3,7–9)], the steady-state levels of each RNA were monitored in the tetO7-ESF1 mutant strain. As seen in Figure Figure5,5, the in vivo levels of these RNAs in the ESF1 mutant were similar to wild type. This suggests that ESF1 is not involved in biogenesis or stability of these snoRNAs.
Using the TAP tag (15), we affinity-purified Esf1p to ascertain which proteins and RNAs were physically associated. Esf1p–TAP co-purified with a number of proteins involved in ribosome biogenesis (Fig. (Fig.6),6), including Nsr1p, Krr1p, Nop1p, Utp22p and Puf6p, all of which are involved in 18S rRNA synthesis (2,5,24–26). In addition, Esf1p–TAP purified with a number of proteins that are components of the ribosome (Rps1p, Rps3p, Rps4p, Rps6p, Rps11p, Rpl2p, Rpl3p, Rpl4p, Rpl7p, Rpl10p, Rpl20p and Rpp0p).
To assay RNAs associated with Esf1p–TAP, RNA was phenol-extracted directly from washed IgG beads. As positive controls, we also affinity-purified Utp18p–TAP, Utp22p–TAP and Nop58p–TAP, which are known to be associated with the U3 snoRNA (2,4,5). Northern blotting revealed that Esf1p–TAP, as well as the three positive controls, co-purified with U3 and U14 snoRNAs (Fig. (Fig.7).7). The protein–RNA associations were stable after washes in 400 mM NaCl. In contrast, TAP-purifications of two other proteins (Krr1p and Nop14p), which are also involved in 18S rRNA biogenesis, did not contain U3 and U14 snoRNAs in the precipitates, in agreement with previous observations (5). Other negative controls (a wild-type strain with no tag and other unrelated proteins Ths1p–TAP, Pus1p–TAP and Gln1p–TAP) also failed to precipitate appreciable amounts of these RNAs. We also detected U14 snoRNA in the Utp18p–TAP and Utp22p–TAP pulldowns, even under very high stringent tandem affinity purifications in which a high-speed centrifugation step was added to the TAP procedure (5), which we note conflicts with previously published observations (4).
We also detected trace amount of 5′-ETS-A0 and 5′-ETS-A1 in the Esf1p–TAP precipitate (Fig. (Fig.7),7), which we have previously shown to associate with components of the SSU processome in the TAP purifications (5). However, we did not detect 20S precursor in the Esf1p immunoprecipitates (data not shown).
We present several lines of evidence that Esf1p is involved directly in 18S rRNA biogenesis. First, depletion of Esf1p led to a virtually complete loss of 20S and 27SA2 precursors, suggesting that the cleavage at site A2 was blocked in the mutant (Fig. (Fig.3).3). In addition, the aberrant 23S RNA species and 35S precursor accumulated in the mutant, indicating inhibition of cleavage events at A0, A1 and A2 (2). Although 18S rRNA biogenesis was affected in the mutant, 25S rRNA levels remained largely normal, suggesting that the 5.8S and 25S rRNAs are formed normally following cleavage at A3 (Fig. (Fig.3).3). Studies of kinetics of rRNA formation in the mutant also suggested that Esf1p mainly affected the biogenesis of 18S rRNA (Fig. (Fig.44).
Among the proteins that co-purified with Esf1p, several are involved in 18S rRNA biogenesis. The nucleolar protein Nop1p is a component of the SSU processome complex (4). Nop1p is also a core component of all Box C/D snoRNPs (2), which include both U3 and U14 snoRNPs. Krr1p is required for 40S ribosome biogenesis (24) and subsequent export to the cytoplasm (5) and was found in the 90S pre-ribosome (5), although in our hands it does not co-purify with U3 or U14 RNAs, consistent with previous results (6) (Fig. (Fig.7).7). Nsr1p, a homolog of mammalian nucleolin (25,26), contains two RRMs (RNA recognition motifs) and a glycine/arginine-rich (GAR) domain. It is required for 18S rRNA synthesis (25,26). Utp22p is also involved in maturation of pre-18S rRNA (5,10) and has been described as a component of the SSU processome (27), although we have previously identified Utp22p in a distinct processome ‘sub-complex’ containing Rrp7p and yeast casein kinase (5). Aside from Utp22p and Nop1p, we did not detect association of Esf1p with any of the components of the SSU processome, indicating that it is not a core component of this large RNP.
Hazbun and colleagues previously identified an Esf1p-associated complex which differs dramatically from ours, containing Bfr2p, Enp2p, Hca4p, Lcp5p, Nop58p and Utp9p (21). Among these proteins, Hca4p, Lcp5p, Nop58p and Utp9p are involved in 18S rRNA synthesis (2,4,28,29). Hca4p is a putative DEAD box RNA helicase and over-expression of HCA4 can suppress a U14 mutant (28). Lcp5p was found to be associated with U3 snoRNA (29) and Nop58p and Utp9p were found in both the SSU processome and 90S pre-ribosome (4,6), although Lcp5p was not found in the SSU processome (4). Bfr2p was also found in the 90S pre-ribosome (6). Hence, Hazbun and colleagues’ data also support the general conclusion that Esf1p is involved in small-subunit biogenesis, although there is a complete lack of overlap with the specific associations we observed.
We propose that Esf1p-containing complexes may be dynamic. Although Esf1p co-purifies with two individual components of the SSU processome (Utp22p and Nop1p) (4,27) (Fig. (Fig.6),6), as well as the U3 and U14 snoRNAs and the 5′-ETS of the rRNA (Fig. (Fig.7),7), all of the associations appear to be sub-stoichiometric, with the possible exception of the ribosomal proteins themselves. This is also clearly the case for the complex described in Hazbun et al. (21). Dynamic complexes might explain why different laboratories obtain quite different results from affinity purifications, since the associated proteins might depend upon the growth phase at harvest, the extraction procedure and the purification protocol used. The presence of Esf1p in multiple pre-ribosomal processing intermediates might also underlie its association with protein components of both the large and small ribosomal subunits. Analysis of mutants in the conserved features of Esf1p (Fig. (Fig.2)2) might help resolve the biochemical role of Esf1p in 18S biogenesis: presumably, they represent discrete domains that mediate physical interactions and/or catalytic functions.
This work was supported by Genome Canada and a CIHR Operating grant to T.R.H.