Dim1p Is Required for Efficient Splicing and Export of mRNA Encoding Lid1p, a Component of the Fission Yeast Anaphase-Promoting Complex

Yeast methods, strains, and media.S. pombe strains used in this study are listed in Table 1. Strains were grown in yeast extract medium or minimal medium with appropriate supplements (21). Crosses were performed on glutamate medium (minimal medium lacking ammonium chloride and containing 0.01 M glutamate [pH 5.6]). Random spore analysis and tetrad analysis were performed as described previously (21). Double mutant strains were constructed and identified by tetrad analysis. Unless otherwise indicated, transformations were performed by electroporation (29). The prp1⁺ gene was tagged at its chromosomal locus to encode a C-terminally tandem affinity purification (TAP)-tagged variant (35) by a PCR-mediated strategy as described previously (2). Proper integration of the TAP cassette was confirmed by PCR and immunoblotting.

Construction of lid1⁺ expression vectors.By using site-directed mutagenesis, NdeI and BamHI restriction sites were placed at the initiating methionine codon and just downstream of the stop codon of lid1⁺ in the genomic clone of lid1⁺ (pKG1295), and the four introns were removed from the lid1⁺ open reading frame (ORF) by site-directed mutagenesis to make pKG1430, a procedure performed by using a Bio-Rad Muta-Gene kit according to manufacturer's instructions, to create the lid1Δi allele. The genomic and cDNA versions of the lid1⁺ ORF as NdeI-BamHI fragments were cloned downstream of the thiamine-repressible nmt1 promoter or its attenuated version, nmt1-41, in the vectors pREP1 (19) and pREP42HA, which add N-terminal hemagglutinin (HA) epitopes (12). All four vectors were able to rescue growth of lid1-6 and the lid1 null mutant. Overexpression of lid1⁺ from these vectors was achieved by growth in the absence of thiamine, while repression was achieved by growth in the presence of 5 μg of thiamine/ml.

To introduce an N-terminal HA tag of lid1Δi at the lid1⁺ genomic locus, an Nde1 fragment encoding three copies of the HA epitope was introduced at the NdeI site of pKG1430, and the lid1⁺ sequences were subcloned into the yeast expression vector pIRT2 that carries the LEU2 selectable marker to make pKG2259. A diploid strain with the relevant genotype lid1⁺/lid1::ura4⁺ leu1-32/leu1-32 was transformed with this vector, and Ura⁺ Leu⁺ colonies were selected and allowed to sporulate. Haploid progeny that were Ura⁺ Leu⁺ were isolated, grown to confluence in the absence of selection, and plated onto appropriate medium containing 5-fluoroorotic acid as described previously (10). Colonies that were Ura⁻ Leu⁻ were then isolated, and the correct replacement of the lid1::ura4⁺ locus with the epitope-tagged version of lid1Δi was confirmed by PCR and Southern blotting.

Plasmids and molecular biological techniques.All plasmid manipulations and bacterial transformations were done according to standard techniques (32). Essential features of plasmid construction are described. All sequencing of plasmid DNA was performed by using Sequenase 2.0 (USB, Cleveland, Ohio) or Thermo Sequenase (Amersham Life Sciences, Cleveland, Ohio) according to the manufacturer's instructions. PCR amplifications were performed by using Taq polymerase and Gene Amp reagents (Perkin-Elmer, Norwalk, Conn.), Pfu polymerase, BioExact (ISC BIOEXPRESS, Kaysville, Utah), or TaqPlus Precision (Stratagene, La Jolla, Calif.) according to the manufacturer's instructions. Amplifications were accomplished by using a PTC-100 programmable thermal controller or a PTC-150 minicycler (MJ Research, Watertown, Mass.).

Immunoprecipitations, immunoblots, and sucrose gradient sedimentation.Protein lysates were made by glass bead disruption of the cell walls in a minimal volume of NP-40 buffer. For denatured lysates, lysed cells were heated to 95°C in sodium dodecyl sulfate (SDS) lysis buffer (10 mM NaPO₄ [pH 7.4], 1.0% SDS, 1 mM dithiothreitol, 1 mM EDTA, 50 mM NaF, 100 μM Na₃VO₄, 4 μg of leupeptin/ml) for 2 min and extracted with NP-40 buffer (6 mM Na₂HPO₄, 4 mM NaH₂PO₄, 1.0% NP-40, 150 mM NaCl, 2 mM EDTA, 50 mM NaF, 100 μM Na₃VO₄, 4 μg of leupeptin/ml) and protease inhibitors. For native lysates, heating in SDS lysis buffer was omitted. For immunoblots, a 1/5 volume of 5× sample buffer was added to the extracts. For quantitative immunoblots, denatured and clarified lysates were normalized by bicinchoninic acid assay (Pierce, Rockford, Ill.) so that equal amounts of protein were loaded into each well of 4 to 20% Tris-glycine polyacrylamide gel. Fractionated proteins were transferred onto Immobilon P membranes (Millipore Corp., Bedford, Mass.) and blotted with anti-Cdc5p (1/5,000), anti-Cdc3p (1/500), and anti-Cdc4p (1/500) rabbit polyclonal antisera and anti-Cdc2p PSTAIR monoclonal antibody (1/5,000) (Sigma, St. Louis, Mo.). 9E10 and 12CA5 mouse monoclonal antibodies (1 μg/ml) were used to detect Myc- and HA-tagged proteins. Goat anti-rabbit and anti-mouse secondary antibodies (Jackson Immunoresearch Laboratories, Inc.) were used at a 1/25,000 dilution. Proteins were visualized with the ECL+ detection system (Amersham) by fluorescence scanning (Storm Phosphoimager; Molecular Dynamics Inc., Sunnyvale, Calif.). ³⁵S-labeled lysates were prepared in an identical manner except that cells were grown overnight in minimal medium and then grown for 4 h in the presence of 1 mCi of ³⁵S-Trans label (ICN Pharmaceuticals, Costa Mesa, Calif.) prior to lysis.

Immunoprecipitations were performed for 1 h on ice followed by a 30-min incubation with 50 μl of a 1:1 slurry of protein A-Sepharose (Pharmacia, Piscataway, N.J.). Immunoprecipitates were washed six times with NP-40 buffer and then resuspended in sample buffer. Anti-Myc immunoprecipitations were performed by using 5 μg of 9E10 antibody and 5 μg of rabbit anti-mouse antibody (Cappel; Orfanon Teknika Corp., West Chester, Pa.). Unless otherwise noted, 12CA5 immunoprecipitations were performed by using 20 μg of 12CA5 which had been coupled to protein A-Sepharose using dimethyl pimelimidate (Sigma). After 1.5 h of incubation, the immunoprecipitates were washed with NP-40 buffer and resuspended in sample buffer.

Proteins were resolved on SDS-6 to 20% polyacrylamide gels. For immunoblotting, proteins were then transferred by electroblotting onto a polyvinylidene difluoride membrane (Immobilon P; Millipore Corp.). Epitope-tagged proteins were detected with 12CA5 (to detect the HA tag) or 9E10 (to detect the Myc tag) antibody at 2 μg/ml in Tris-buffered saline followed by alkaline phosphatase-conjugated goat anti-mouse polyclonal antibodies (Sigma). Cdc2p was detected with anti-PSTAIRE antibodies (immunoblots were visualized by using enhanced chemiluminescence [ECL; Amersham]). For visualization of ³⁵S-labeled proteins, the protein gels were fixed, treated for fluorography (Amplify; Amersham), dried, and exposed to film.

Protein complexes were obtained by using the TAP strategy as described previously (35), except that the lysates were clarified at 3,000 rpm on a tabletop GS-6R centrifuge in lieu of ultracentrifugation. TAP pellets were subjected to mass spectrometric analyses as described previously (8). Sucrose gradient sedimentation analysis was performed exactly as described previously (20), except that gradients were centrifuged for 16 h at 25,000 rpm.

RNA and Northern blots.Total RNA from S. pombe cells was prepared as described previously by Moreno et al. (21). The S. cerevisiae dib1Δ shutoff strain (KLG1806) was grown under permissive conditions and shifted to restrictive conditions as described previously (5). The control mutants used were prp3-1 (ts125) (KLG1825) (7, 39), prp18 (ts503) (KLG1229) (39), and cdc28-1N (KLG1760) (25). Total RNA was prepared from these cells by extraction with hot acidic phenol as described previously (11). To detect mRNAs, total RNA (20 μg) was resolved with formaldehyde agarose gels and capillary blotted to GeneScreen⁺ (Dupont-NEN, Boston, Mass.) or Duralon-UV (Stratagene). tf2d RNA was detected by using ³²P-labeled oligonucleotides complementary to both intronic (TFIID I) and exonic (TFIID E) sequences as described previously (17, 26). his3⁺ RNAs were detected by using the ³²P-labeled EcoRV-DraI segment of the genomic clone (10) as a probe. Blots of S. cerevisiae RNAs were hybridized with labeled probes from RP51a, DYN2, and GLC7 ORFs or a PCR fragment representing the ACT1 intron sequences. Blots were exposed to PhosphorImager screens and visualized by using MD Image Quant software version 3.3 (Molecular Dynamics).

Microscopy.All microscopy was performed with a Zeiss Axioskop II equipped with a z-focus motor drive, and images were captured with an Orca II charge-coupled-device camera (Hamamatsu, Japan). Images were obtained, processed, and analyzed with OpenLab 2.1.3 software (Improvision, Lexington, Mass.).

In situ hybridization.Cells were fixed in suspension with 3.7% formaldehyde for 30 min, washed two times in 0.1 M potassium phosphate, pH 6.5 (K-Pi buffer), washed once in K-Pi buffer plus 1.2 M sorbitol (K-Pi/SORB), and resuspended in 1 ml of K-Pi/SORB. Three microliters of β-mercaptoethanol was added, and cells were incubated for a further 10 min. Thirty microliters of Zymolase 20T (10 mg/ml) was then added to cells, and they were incubated for 30 to 60 min with rotation. Cells were then washed three times with K-Pi/SORB, once with K-Pi, once with K-Pi plus 0.1% NP-40, and once with K-Pi. The cells were then resuspended in a solution containing 100 μl of 50% formamyde, 4× SSC (1× SSC is 0.15 M NaCl plus 0.015 M sodium citrate), 1× Denhardt's solution, 125 μg of tRNA/ml, 10% dextran sulfate, and 500 mg of denatured salmon sperm DNA/ml and incubated for 1 h at 37°C. A 418-pg/ml oligo(dT)₅₀ probe, 3′-end labeled with digoxigenin-11-dUTP (Boehringer Mannheim) as previously described (13), was added to the cells, and they were incubated overnight at 37°C while rotating. The cells were then washed for 1 h in 2× SSC at room temperature, 1 h in 1× SSC at room temperature, 30 min in 0.5× SSC at 37°C, 30 min in 0.5× SSC at room temperature, 5 min in phosphate-buffered saline containing 1% bovine serum albumin (PBAL) at room temperature, and 1 h in PBAL at room temperature. Cells were resuspended in 50 μl of PBAL, and rabbit polyclonal anti-digoxigenin antibody conjugated to fluorescein isothiocyanate was added at a dilution of 1:25 and then incubated for 3 to 4 h at room temperature. Cells were subsequently washed two to three times in PBAL and mounted onto slides as described previously (13).

Yeast two-hybrid analysis.The yeast two-hybrid system used in this study was described previously (15). The indicated cDNAs were cloned into the bait plasmid pGBT9 and/or the prey plasmid pGAD424 (Clontech, Palo Alto, Calif.) and sequenced to ensure the absence of PCR-induced mutations and to ensure that the correct reading frame had been retained. To test for protein interactions, both bait and prey plasmids were cotransformed into S. cerevisiae strain PJ69-4A.β -Galactosidase reporter enzyme activity in the two-hybrid strains was measured by using the Galacto-Star chemiluminescent reporter assay system according to the manufacturer's instructions (Tropix Inc., Bedford, Mass.), with the exception that cells were lysed by glass bead disruption. Each sample was measured in triplicate. Reporter assays were recorded on the Mediators PhL luminometer (Aureon Biosystems, Vienna, Austria).

Green RNA.The system for imaging the localization of specific mRNA transcripts was adapted from a similar system in S. cerevisiae (a generous gift of Kerry Bloom [3]). The green fluorescent protein (GFP)-fused MS2 coat protein (CP) (CP-GFP) was subcloned by PCR from pCP-GFP into the NdeI/NotI sites of pREP41NT (35), creating the pREP41CP-GFP vector. Primers used to amplify CP-GFP were pCP-GFP-For (5′-TAGGCGCGCCCATATGGCTTCTAACTTTACTCAGTTCGTTCTCGTCG-3′) and pCP-GFP-Rev (5′-TTTCCTTTTGCGGCCGCCCGGGTCGACTTATTTGTATAGTTCATTG-3′). To create a host vector for RNA transcripts of interest, an attenuated adh1 promoter (a generous gift of Charlie Albright) was subcloned from pSK utilizing PstI/XhoI and ligated into the pREP2 vector (19) previously digested with PstI/SalI, creating the pRAM vector (pREP with adh mutated). The 154-bp MS2 binding site sequence was excised from pIIIA/MS2-2 by digestion with EcoRI. This fragment contains two tandem copies of the 25-nucleotide MS2 coat protein binding site and a single adjacent SmaI site for cloning sequences of interest. The fragment was first treated with Klenow fragment to create blunt ends and subsequently ligated into the SmaI site of pRAM to create pRAM-MS2. All sequences examined were cloned into the SmaI site of pRAM-MS2, and visualization was accomplished by cotransformation with pREP41CP-GFP and examination of GFP fluorescence in live cells of the strains indicated.

dim1⁺ function is required for pre-mRNA splicing.To learn more about the biochemical function of Dim1p, we examined the biochemical basis for the decreased abundance of the APC/C component Lid1p in the absence of dim1⁺ function. Because Lid1p is involved in the process of proteolysis, we began our analysis by examining Lid1p degradation rates in the presence and absence of dim1⁺ function. For this purpose, the lid1⁺ ORF including its four introns was tagged with sequences encoding three copies of the HA epitope at its 5′ end, and the fusion construct was placed under control of the thiamine-repressible attenuated nmt1 promoter (nmt41). The plasmid expressing HAlid1⁺ was able to rescue both the lid1-6 and lid1 null alleles, and overexpression elicited no obvious phenotype in any background (data not shown). HAlid1⁺ was then overexpressed in wild-type and dim1-35 cells. Although HA-Lid1p was readily detected in lysates prepared from wild-type cells, it was present at significantly reduced levels in dim1-35 cells even when the cells were grown at 25°C, a temperature fully permissive for growth (Fig. 1A). This result reproduces what was observed previously for endogenous Lid1p in the absence of Dim1p function (4).

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FIG. 1.

HA-Lid1p cannot be overproduced in the absence of dim1⁺ function. Wild-type and dim1-35 cells were transformed with pREP42HAlid1⁺. Transformants were grown at 25°C in the absence of thiamine for 20 h, and samples were collected. A) Total protein lysates were prepared from the samples and resolved by SDS-polyacrylamide gel electrophoresis (PAGE). Following immunoblotting with 12CA5 to detect HA-Lid1p and anti-Arp3p serum (19a) to detect Arp3p as a loading control, proteins were visualized by enhanced chemiluminescence. B) After maximal induction of pREP42HAlid1⁺ in wild-type (upper panel) and dim1-35 (lower panel) cells, 5 μg of thiamine/ml was added to the medium to prevent further expression from the nmt41 promoter. Equal numbers of cells were collected at hourly intervals, and protein lysates were prepared. Proteins were resolved by SDS-PAGE. HA-Lid1p was detected with 12CA5 antibodies, and Cdc2p, which served as a loading control, was detected with anti-PSTAIR monoclonal antibody. The immunoblot in the lower panel was developed for longer than the immunoblot in the upper panel so that the lower level of HA-Lid1p in dim1-35 cells could be visualized throughout the course of the experiment.

Despite the lower level of total HA-Lid1p in dim1-35 cells compared to wild-type cells, we were able to determine the relative half-life of HA-Lid1p in the two strains. After maximal expression was achieved by growing cells in the absence of thiamine, synthesis of the RNA and protein was repressed by adding thiamine and cycloheximide, respectively, to the medium. The amount of HA-Lid1p that remained was determined at hourly time points by immunoblotting with antibodies to the HA epitope. In wild-type cells, after an initial burst of protein production, the amount of HA-Lid1p declined steadily over the course of the experiment (Fig. 1B, upper panel). Although expressed at significantly reduced levels from the onset, the half-life of HA-Lid1p in dim1-35 cells appeared even longer than that in wild-type cells (Fig. 1B, lower panel). Thus, increased rates of HA-Lid1p protein degradation could not explain the decreased abundance of Lid1p in the absence of dim1⁺ function.

Based on this result, we then predicted that HA-Lid1p was synthesized at reduced levels in the absence of dim1⁺ function. To test this hypothesis, wild-type and dim1-35 cells expressing maximal levels of HA-Lid1p were pulsed for 10 min with ³⁵S-Trans label, and the amount of HA-Lid1p produced was determined by immunoprecipitation. Despite similar amounts of ³⁵S incorporation into the strains during a 10-min pulse, there was significantly less HA-Lid1p produced in dim1-35 cells than that produced in wild-type cells (data not shown).

We next examined the steady-state level of HAlid1⁺ mRNA produced from the nmt41 promoter in dim1-35 cells relative to wild-type cells by Northern blot analysis. Although the total levels of nmt41HAlid1⁺ RNA production were similar at both permissive and nonpermissive temperatures, the nmt41HAlid1⁺ RNA was split into two bands in the dim1-35 cells (Fig. 2A, strain 3). The faster-migrating band comigrated with the nmt41HAlid1⁺ RNA from wild-type cells. Since lid1⁺ contains four introns, it seemed likely that the upper band represented an unspliced form of nmt41HAlid1⁺ RNA. To test this possibility directly, the four introns were removed from the nmt41HAlid1⁺ construct. When the cDNA version of lid1⁺ was overexpressed in dim1-35 cells in parallel with the intron-containing form, the upper band was no longer observed (Fig. 2A, strain 5). Thus, dim1-35 cells appeared to be defective in the splicing of lid1⁺ pre-mRNA.

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FIG. 2.

Cells lacking dim1⁺ function are defective for pre-mRNA splicing. A) The lid1⁺ ORF with (strains 3 and 4) or without (strains 5 and 6) its four introns was introduced into the pREP42HA vector. Wild-type cells (strains 2, 4, and 6) and dim1-35 cells (strains 1, 3, and 5) were transformed with vector alone (strains 1 and 2) or the pREP42HAlid1⁺ constructs (strains 3 to 6). Transformants were grown at 25°C in the absence of thiamine for 18 h and then transferred to 36°C for 0 or 4 h. Total RNA was prepared and subjected to Northern analysis with a fragment of the lid1⁺ ORF as probe. Note by the absence of bands in 1 and 2 that endogenous levels of lid1⁺ RNA are not detected in these exposures. B) Total RNA was purified from wild-type (wt), prp2-1 shifted to the nonpermissive temperature for the indicated number of hours, a strain containing dim1::his3⁺ and nmt1-T81 dim1⁺ integrated at the leu1 locus grown in presence of thiamine for the indicated number of hours, and dim1-35 cells shifted to the nonpermissive temperature for the indicated number of hours. Twenty micrograms of total RNA from each sample was resolved by electrophoresis and subjected to Northern blot analysis with oligonucleotide probes complementary to the intron and exon sequences within the tf2d gene. his3⁺ RNAs were detected with a ³²P-labeled probe derived from the genomic clone. PC, precursor mRNA, M, mature mRNA. C) S. cerevisiae cells lacking DIB1 are defective in pre-mRNA splicing. Strain KGY1023 was maintained in synthetic medium containing raffinose and galactose. DIM1 expression was repressed by shifting the cells to synthetic medium containing glucose (SD). Aliquots of cells were collected at the number of hours indicated following the shift into synthetic medium containing glucose. Total RNA was also purified from temperature-sensitive mutants prp3-1, prp18 (ts503), and cdc28-1N shifted to the restrictive temperature (35.5°C) for the number of hours indicated. Twenty micrograms of total RNA was electrophoresed and blotted. Northern blots probed with the ACT1 intron sequence or the DYN2, GLC7, and RP51a ORFs. Note the mature form of GLC7 mRNA does not decline because of the presumed longer half-life of the species.

The failure of dim1-35 cells to splice lid1⁺ mRNA might explain the reduced levels of Lid1p in this strain. To test whether Dim1p was required for general pre-mRNA splicing in vivo, RNA was prepared from wild-type cells, prp2-1 (a bona fide pre-mRNA splicing mutant) cells, cells genetically depleted for dim1⁺ (a dim1 null mutant containing nmt1-regulatable dim1⁺), and dim1-35 cells, and the RNA was subjected to northern analysis using probes directed at two intron-containing genes, tf2d and his3⁺. Like prp2-1 cells, cells lacking dim1 function accumulated unspliced RNAs (Fig. 2B).

We next tested whether the S. cerevisiae ortholog of Dim1p, termed Dib1p, was required for pre-mRNA splicing, since Dib1p had also been implicated in pre-mRNA splicing due to its copurification with the U4/U6.U5 tri-snRNP (14, 34). Previously, in order to investigate the phenotype of cells lacking Dib1p function, we had created a conditional expression strain, KGY1023 (5). Because GAL1-driven DIB1 was not sufficiently repressed by glucose addition, we made use of the ubiquitin-N-degron tagging strategy described previously by Althoefer et al. (1). A plasmid expressing budding yeast DIB1 still allowed growth in the presence of glucose. However, a single integrated copy of GALS::UBdim1⁺ rescued growth under inducing conditions but failed to rescue growth under repressing conditions. Both S. pombe dim1⁺ and mouse mDim1 rescue the dib1 null mutation (5). KGY1023, which lacks endogenous DIB1 and harbors plasmid-borne S. pombe dim1 cDNA under control of the GAL1 promoter, arrests growth following 6 h of glucose repression (5). KGY1023 mRNA was compared to that isolated from three control strains: (i) prp3-1, a positive control for a defect in the first step of splicing (39); (ii) prp18 (ts503), a positive control for a defect in the second step of splicing (39); and (iii) cdc28-1N, a G₂ arrest (25) control to ensure that any observed splicing defects were not secondary to cell cycle arrest. We assayed four intron-containing transcripts: ACT1 and RP51a, which are routinely used to analyze splicing defects in prp mutants; DYN2, a transcript in S. cerevisiae that contains two introns; and GLC7, which encodes a cell cycle-regulated protein. When dim1 expression was repressed, intron-containing forms of all four transcripts steadily accumulated to levels comparable to what was observed in the prp mutants (Fig. 2C). Also, the levels of mature DYN2 and RP51a decreased throughout the time course. These results were comparable to those observed with the prp3-1 mutant but distinct from those observed with prp18 and cdc28-1N. These data therefore suggest that DIB1 is essential, either directly or indirectly, for the first step of pre-mRNA splicing in vivo.

Dim1p copurifies with known splicing factors.To determine whether Dim1p was a part of the S. pombe U4/U6.U5 tri-snRNP, similar to Dib1p in S. cerevisiae (14, 34), we examined whether it was present in a high-molecular-weight complex by sucrose gradient sedimentation. We found that a percentage of Dim1p sedimented deeper into the gradient than Cdc5p (which runs at approximately 40S) (20) (Fig. 3A), indicating that Dim1p was present in a complex considerably larger than what would be expected from the tri-snRNP (34). The remainder of Dim1p sedimented near the top of the gradient, consistent with a monomer or small complex (Fig. 3A). To determine if other components of the S. pombe U4/U6.U5 tri-snRNP behaved similarly on sucrose gradients, we modified the S. pombe ortholog of S. cerevisiae PRP6, prp1⁺ (also known as zer1⁺) (38), at its endogenous locus to encode C-terminally Myc13- or TAP-tagged versions of Prp1p (30, 35). Both tagged strains grew normally, suggesting that the epitope did not compromise the function of Prp1p. By sucrose gradient fractionation, Prp1p-Myc13 sedimented deep into the gradient similarly to one portion of Dim1p. To determine if the complex that contained Prp1p also contained Dim1p, tandem affinity purification was carried out on two separate occasions from a prp1-TAP strain, and the protein composition of a portion of each TAP complex was analyzed by silver staining (Fig. 3B), with the remainder analyzed by multidimensional tandem mass spectrometry (16). Proteins identified from both purifications that were absent from TAP purifications performed on untagged cells or from unrelated TAP purifications (data not shown) are listed in Table 2. As a means of comparison, results from the purification of the S. cerevisiae penta-snRNP complex (34) and the S. pombe Cdc5p splicing complex (23) are also shown. In both Prp1p-TAP purifications, Dim1p was present, as were most components of the U4/U6.U5 tri-snRNP and the U2 snRNP. Indeed, the protein composition of this complex is very similar to the recently described human BΔ1 complex that lacks the U1 snRNP and the nineteen complex (18). The amino acid sequence coverage of Prp1p was 60%, and the greatest sequence coverage of copurifying proteins was obtained for Dim1p and Prp31p, at 58 and 61%, respectively. While these results are not quantitative, they did raise the possibility that Dim1p interacted with Prp1p directly or indirectly through an association with Prp31p, and we tested whether Dim1p was capable of binding either of these two proteins. We found that Dim1p interacted with Prp1p by two-hybrid analysis using full-length constructs of each protein, but it did not show an interaction with Prp31p (Fig. 3C). In contrast, Prp1p showed a strong interaction with both Dim1p and Prp31p (Fig. 3C and D), indicating that Prp1p might bind both proteins directly and simultaneously.

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FIG. 3.

Dim1p copurifies with known splicing factors. (A) Protein lysates of dim1-HA or prp1-myc strains were fractionated by sucrose gradient sedimentation. Fractions were collected from the bottom (fraction 1) of the gradients, resolved by SDS-PAGE, and then immunoblotted with the 12CA5 or 9E10 antibody to detect Dim1p-HA (upper panel) and Prp1p-Myc (lower panel), respectively. The signal on the right-hand portion of the anti-Myc blot is a nonspecific blotch. The positions of sedimentation markers are provided. (B) A silver-stained gel of the purified Prp1p-TAP complex. (C and D) The indicated proteins (bait vector/prey vector) were tested by two-hybrid analysis. LEU⁺ TRP⁺ transformants were tested for growth on selective medium (data not shown) and assayed for β-galactosidase activity measured in relative light units.

The dim1-35 mutation selectively affects the production of Lid1p.A block to pre-mRNA splicing as in the dim1-35 mutant would be expected to affect the levels of most transcripts and their protein products. However, this seemed incongruous for two reasons. First, we had used Cdc2p and Arp3p as loading controls for our immunoblots and observed no difference in their abundance between wild-type and dim1-35 cells (4) (Fig. 1A), although both the cdc2⁺ and arp3⁺ primary transcripts contain four introns. Second, it would be difficult to imagine a scenario in which a general block in protein production would generate the dim1-35 mutant phenotype. To examine this question more carefully, we compared the levels of several proteins whose pre-mRNAs contain introns in the dim1-35 mutant and wild-type cells. The levels of Cdc3p (profilin), Cdc4p (a myosin light chain), Cdc5p (a pre-mRNA splicing factor), Arp3p (a component of the Arp2/3 F-actin-nucleating complex), Cwf2p (a pre-mRNA splicing factor), and Cwf3p (a pre-mRNA splicing factor) were not significantly different in the dim1-35 mutant relative to wild-type cells using the amount of Cdc2p in the lysates as a loading control (Fig. 4A). Only Lid1p levels were found to be significantly different (Fig. 4A). This was also true if the specific signals were quantitated against total protein loaded onto the gels rather than Cdc2p abundance (data not shown). Thus, the dim1-35 mutation selectively affects the production of Lid1p among tested proteins.

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FIG. 4.

Dim1p affects Lid1p levels independently of pre-mRNA splicing. (A) Cells were grown at permissive temperature (25°C) and shifted to 36°C for 4 h. Protein lysates were prepared and normalized according to protein concentration. Equal amounts of protein from wild-type (wt) (KGY246) and dim1-35 (KGY390) cells were blotted with specific antisera for Cdc5p, Cdc3p, Arp3p, Cdc4p, and Cdc2p or from strains that express Myc or HA epitope-tagged proteins (KGY1365, KGY1739, KGY1302, KGY1305, KGY1420, KGY3211, KGY1430, and KGY3210) with 9E10, 12CA5, and Cdc2p antibodies. Immunoblots were analyzed with a Molecular Dynamics Storm PhosphorImager, and the intensity of each band was normalized against the Cdc2p loading control. Results are presented as means ± standard deviations (n = 3 to 10). (B)Wild-type (lanes 1, 3, and 5) and dim1-35 (lanes 2, 4, and 6) cells were transformed with empty plasmid (lanes 1 and 2), the pREP42HAlid1⁺ construct containing introns (lanes 3 and 4), or the pREP42HAlid1⁺ construct lacking introns (lanes 5 and 6). Following maximal RNA production at 20 h of growth in the absence of thiamine and a shift to 36°C for 4 h (Fig. 2), protein lysates were prepared and resolved by SDS-PAGE. HA-Lid1p (upper panel) and Cdc2p which served as a loading control (lower panel) were detected by immunoblot analysis. (C) Lysates were prepared from wild-type cells (lane 1) or wild-type and dim1-35 cells expressing HA-lid1Δi from the endogenous lid1 promoter (lanes 2 and 3, respectively) after a shift to 36°C for 4 h. Following separation by SDS-PAGE, HA-Lid1p (upper panel) and Cdc2p as a loading control (lower panel) were detected by immunoblotting.

If Dim1p controlled the levels of Lid1p solely by regulating the splicing of lid1⁺ RNA, then production of Lid1p in dim1-35 cells should be restored to wild-type levels by the removal of the four introns from the lid1⁺ ORF. To test this hypothesis, we examined the level of HA-Lid1p produced from the lid1⁺ cDNA (lid1Δi) under control of the nmt41 promoter. Unexpectedly, we found that HA-Lid1p levels were still reduced in dim1-35 cells relative to that of wild-type cells, although the level of Cdc2p did not vary (Fig. 4B, lanes 5 and 6). We then considered the possibility that overproduction of HA-Lid1p from a heterologous promoter was overwhelming the capacity of dim1-35 cells to produce HA-Lid1p. Therefore, we constructed a gene replacement strain. First, we introduced sequences encoding three copies of the HA epitope at the 5′ end of the open reading frame of the intron-deleted version of lid1⁺. Next, the tagged version of lidΔi was used to replace the endogenous gene (see Materials and Methods) so that expression would occur from the endogenous lid1 promoter. The HAlid1Δi strain was wild type in morphology and growth rate (data not shown). The HAlid1Δi allele was then combined with the dim1-35 mutation, and endogenous HA-Lid1p levels were examined after a shift to a restrictive temperature. The amount of HA-Lid1p produced in dim1-35 cells was barely detectable and significantly less than that in wild-type cells (Fig. 4C). Thus, there appears to be a second block to Lid1p production downstream of pre-mRNA splicing in dim1-35 cells.

Dim1p function is required for efficient pre-mRNA export.The process we thought to examine next in cells lacking dim1⁺ function was the export of RNA from the nucleus. The localization of poly(A)⁺ RNA was examined in dim1-35 and dim1Δ cells and compared with that in wild-type cells and a bona fide nuclear export mutant, rae1-1 (9). In wild-type cells, poly(A)⁺ RNA was not detected in the nucleus at appreciable levels (Fig. 5A). In the dim1-35 mutant, poly(A)⁺ RNA could be detected in the nucleus even at permissive temperature, and staining within the nucleus increased during the temperature shift (Fig. 5A). However, nuclear accumulation was neither as complete nor as rapid as that observed in rae1-1 cells (Fig. 5A). In the dim1Δ cells maintained by dim1⁺ expressed from a regulatable thiamine-repressible promoter, nuclear pre-mRNA accumulation was observed concomitantly with the timing of promoter repression (Fig. 5B). Furthermore, this accumulation paralleled the timing of the loss of pre-mRNA splicing (Fig. 5C).

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FIG. 5.

Localization of poly(A)⁺ RNA in dim1-35 cells. (A) poly(A)⁺ RNA was detected by fluorescence in situ hybridization in wild-type, rae1-1, and dim1-35 cells grown at 25°C and shifted to 36°C for the times indicated. (B and C) dim1::his3⁺ cells carrying a single integrated copy of nmt81::mDIM1⁺ (KGY1180) were grown in minimal medium lacking thiamine and then shifted to medium containing thiamine for the times indicated. DAPI, 4′,6′-diamidino-2-phenylindole. (B) poly(A)⁺ RNA was detected by fluorescence in situ hybridization. (C) Total RNA from these cells was examined by Northern blot analysis for the accumulation of TFIID pre-mRNA. PC, precursor; M, mature RNA.

The dim1-35 mutation selectively affects the export of lid1 mRNA.One possible explanation for the selective effects of Dim1p loss of function on Lid1p levels is that the function of Dim1p in either pre-mRNA splicing and/or nuclear mRNA export is specific to lid1 transcripts or a subset of transcripts that includes lid1. Since our data indicated a generalized defect of pre-mRNA splicing (Fig. 2), we examined the specificity of the dim1-35 mRNA export defect for lid1 transcripts. To address this, we adapted the S. cerevisiae “green RNA” system, developed for live cell monitoring of specific mRNA transcripts (3, 33), for use in S. pombe (see Materials and Methods). In wild-type cells, the lid1⁺ GFP-labeled mRNA transcript (gRNA) was not detected in the nucleus at appreciable levels (Fig. 6). However, in the dim1-35 mutant, lid1⁺ gRNA could be detected in the nucleus at the restrictive temperature (Fig. 6). However, as observed with total mRNA (Fig. 5), nuclear accumulation was not as complete or as rapid as that observed in rae1-1 cells (Fig. 6). The accumulation of lid1 transcript in dim1-35 is unrelated to the role of Dim1p in splicing, as both wild-type and lid1Δi transcripts showed comparable levels of accumulation. Furthermore, no appreciable accumulation of lid1 transcript was observed in prp2-1 cells, which are strongly inhibited for pre-mRNA splicing, at the restrictive temperature (data not shown). To investigate the specificity of the RNA export defect, we also examined the localization of the unrelated wsp1 transcript. While wsp1 transcripts with or without introns do accumulate in the nucleus of rae1-1 cells (Fig. 6 and data not shown), which show a block of generalized mRNA export (9), no significant accumulation of these transcripts was observed in dim1-35 cells (Fig. 6).

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FIG. 6.

Localization of individual transcripts in dim1-35 cells. lid1 without (A to D) (lid1 cRNA) or with (E to H) (lid1 gRNA) introns as well as control wsp1 gRNA (I to L) were generated by placing them downstream of the MS2-CP binding sites in pRAM-MS2. Coexpression of the gRNA expression constructs with pREP41CP-GFP allowed visualization of their localization at either 25°C (A, E, and I) or 36°C (B to D, F to H, J, and K) in dim1-35 cells (A, B, E, F, I, and J), rae1-1 cells (C, G, and K), and wild-type (wt) cells (D, H, and L).