Purification and structure determination of theS. cerevisiae Cet1-Ceg1 complex

A Cet1 polypeptide encompassing amino acids 241–549 suffices for triphosphatase activity in vitro and for mRNA capping in vivo (Lehman et al., 1999).S. cerevisiae triphosphatase Cet1 (241–549; referred to as Cet1 hereinafter) and full length guanylyltransferase (Ceg1 1–459) were co-expressed inE. coli (Experimental Procedure). Although Ceg1 was the only protein fused to a His₆-Smt3 tag, Cet1 and Ceg1 co-purified by metal affinity chromatography. After removal of His₆-Smt3 tag by digestion with the Smt3 protease Ulp1 (Mossessova and Lima, 2000), Ceg1 and Cet1 retained the ability to interact as evidenced by co-elution during anion-exchange and gel filtration chromatography (Figures S1A and S1B). Cet1-Ceg1 eluted in two peaks during anion exchange chromatography and analysis of these peaks revealed that one contained a 2:1 complex between Cet1 and Ceg1 (peak 1) while the other contained a 2:2 complex (peak 2) (Figure S1C). Crystals were obtained after a few days for peak fractions containing 2:2 Cet1-Ceg1 while those containing 2:1 Cet1-Ceg1 took weeks to form crystals. Crystals from either preparation were isomorphous suggesting that both contained complexes of similar composition. Based on the time it took to obtain crystals for the 2:1 complex, and based on our structure of the Cet1-Ceg1 complex (see below), we infer that the 2:1 complex was in equilibrium with the 2:2 complex and it was the 2:2 complex that crystallized. A complete data set was collected from a single crystal to a resolution of 3 Å and experimental phases were obtained from complete data sets collected from two crystals that were derivatized with thimerosal for 8 or 16 hours, respectively. A complete data set was obtained from a crystal containing selenomethionine substituted proteins at 4.3 Å resolution (Hendrickson et al., 1990) (Table 1) and used to confirm positions of methionine in our model.

Native and derivative data sets were initially reduced in space group P6₃22 and used to calculate phases. An atomic model for Ceg1 was manually built into electron density and one Cet1 protomer was docked into the experimental electron density based on a model derived from previous Cet1 structures (Lima et al., 1999). Inspection of experimental electron density revealed additional electron density consistent with another molecule of Cet1 that was intertwined with the one that was docked into the density map (Figure S2A and S2C). To confirm our model and positions for Cet1 and Ceg1 in the asymmetric unit, a complete data set was collected at a single wavelength from a crystal containing selenomethionine substituted proteins (Hendrickson et al., 1990). An anomalous difference Fourier map revealed electron density proximal to many of the positions for methionine side chains thus confirming our model (Figure S2B).

Assuming static disorder in the lattice, we attempted to refine a model encompassing one Ceg1 molecule and two overlapping Cet1 protomers in space group P6₃22, as well as two Ceg1 molecules and two overlapping Cet1 homodimers in space groups P321, and P6₃, but this approach failed to provide an adequate physical explanation for the two overlapping molecules of Cet1 (See Methods;Figure S2). Subsequent statistical analysis of the data revealed that Cet1-Ceg1 crystals were perfectly twinned (Padilla and Yeates, 2003) and analysis of a model in space group P6₃ provided a physical explanation of how the complex is twinned in the lattice (See Methods;Figure S3). Native and derivative data sets were reprocessed in space group P6₃ and experimental phases calculated. The resulting experimental electron density was used to evaluate and rebuild the models for one Cet1 homodimer and two molecules of Ceg1 in the asymmetric unit (Figures S2A and S2B). The final model was refined by applying the twin operator during refinement with tight NCS restraints for backbone atoms using Refmac (Murshudov et al., 1997) resulting in a R_work and R_free of 0.249 and 0.298, respectively (Table 1;Figure S4).

Overall organization of the Cet1-Ceg1 complex and structure of Cet1

Central to the Cet1-Ceg1 complex is one Cet1 homodimer that is bordered on each side by a molecule of Ceg1 (Figure 1). The two Cet1 protomers form an extensive dimer interface that is nearly identical to that observed in structures of Cet1 alone (Figures 2A and 2B;Lima et al., 1999). Interactions between the respective Cet1 and Ceg1 protomers primarily involve contacts between the Ceg1 OB domain and an extended Cet1 element located near the N-terminus of our construct that originates from the distal Cet1 protomer, thus resulting in a swapped configuration for the respective N-termini (Figure 1 andFigure 2).

The yeast RNA triphosphatase Cet1 includes a signature tunnel composed of eight anti-parallel β-strands which direct several catalytic amino acid side chains into the tunnel interior (Figures 2C and 2D;Lima et al., 1999). In this previous work, Cet1 was crystallized in the presence of ammonium sulfate and manganese ions. The respective positions of these ligands suggested that the sulfate ion might mimic the position of the nascent mRNA γ-phosphate since it was observed coordinated by several basic side chains and the manganese ion, the latter was coordinated by several conserved acidic residues that emanate from the bottom of the tunnel (Figure 2C;Lima et al., 1999). No additional density was observed in the present structure adjacent to the aforementioned active site residues within the tunnel, thus we infer that both active sites are devoid of ligands (Figure 2D).

Cet1 amino acids 268–539 adopt similar conformations in the Cet1-Ceg1 complex when compared to Cet1 structures alone (Lima et al., 1999), although major differences were observed with respect to the conformation of Cet1 amino acids 241–261. In previously determined Cet1 structures, residues 241–261 adopted an extended conformation across the Cet1 dimer interface forming contacts to both Cet1 protomers (Figure 2A) and the functional importance of these contacts was highlighted by a Cet1 construct that contained an N-terminal deletion up to amino acid 275 which resulted in a catalytically active, albeit monomeric form of Cet1 (Lehman et al., 1999). Amino acids 268 to 275 adopt similar conformations to that observed in previous Cet1 structures. Cet1 amino acids 261 and 268 exhibited very weak electron density and were presumed disordered while amino acids 241 to 261 adopt a distinctly different conformation within the context of interactions with the Ceg1 OB domain (Figure 2D andFigure 3). The disordered linker between amino acids 261 and 268 is consistent with previous studies that showed this region was highly susceptible to protease digestion in solution (Lehman et al., 1999).

The structure of Ceg1 in the Cet1-Ceg1 complex

The Ceg1 guanylyltransferase molecules in the Cet1-Ceg1 complex adopt similar open conformations with respect to the oligonucleotide binding (OB) and nucleotidyl transferase (NT) domains (Figure 3). Examination of the Ceg1 active site did not reveal electron density for either substrate (GTP) or the product lysyl-GMP. Comparison to the only other structure of a cellular guanylyltransferase (C. albicans guanylyltransferase Cgt1) shows that the Ceg1 OB domain swings an additional 20° away from the NT domain in comparison to that observed for the open configuration of Cgt1 (Figure 4;Fabrega et al., 2003). Guanylyltransferase capping enzymes undergo conformational changes that involve opening and closing of the OB and NT domains to facilitate metal-dependent transfer of GMP from GTP to the active site lysine in step 1 and transfer of the GMP to the diphosphate terminated 5’ end of nascent mRNA in step 2 (Shuman and Lima, 2004). Comparison of Ceg1 in the open conformation to the Chlorella virus guanylyltransferase in the closed conformation (Hakansson et al., 1997;Hakansson and Wigley, 1998) illustrates the extent of the conformational changes and that the Ceg1 OB domain would be required to rotate 60° to achieve the closed conformation as calculated by Dyndom (Hayward and Berendsen, 1998).

Each of the Ceg1 OB domains engage Cet1 amino acids 245–261 in a similar manner (Figure 5), although the two Ceg1 molecules were observed in distinct orientations with respect to the adjacent Cet1 dimer (Figure 3). In one Cet1-Ceg1 interface, the OB domain interacts with Cet1 (2400 Ų total buried surface area) and in this conformation the majority of buried surface area (1900 Ų) is derived from contacts between the OB domain and Cet1 amino acids 245–261 although a few minor contacts were observed between a Cet1 β-strand (aa 368–472) and the OB domain (Figure 3B). In this configuration the Ceg1 NT domain makes no contacts to Cet1 and the OB and NT domains appear free to open and close to facilitate GMP transfer. Similar contacts were observed in the other Cet1-Ceg1 complex but in this instance Ceg1 NT domain makes a few contacts to Cet1 that would be predicted to block Ceg1 from closing to promote GMP transfer (Figure 3A). Because this interface buries only 200 Ų of additional total surface area, we believe the Ceg1 NT domain would remain free to swing away from this contact in solution during capping.

Interactions between Cet1 and the Ceg1 OB domain and functional analysis in vivo

Interactions between Cet1 and Ceg1 are dominated by contacts between Cet1 amino acids 245–261 and the Ceg1 OB-fold domain which bury 1900 Ų of total surface area in the respective interface (Figures 5A and 5B) and this Cet1 element includes the WAQKW amino acid motif which has been shown to be an essential for function in vivo and for Cet1-Ceg1 interaction in vitro (Ho et al., 1999;Lehman et al., 1999;Takase et al., 2000). The extended conformation observed for Cet1 amino acids 245–261 enables many of the Cet1 side chains to interact directly with the Ceg1 OB surface which is lined by several aliphatic side chains (Figure 6A) that are conserved betweenS. cerevisiae Ceg1 andC. albicans Cgt1 (Figure 6B).

Cet1 Pro245 and Ile246 pack on top of Trp247 whose side chain makes van der Waals contacts to Ceg1 Trp337, Leu340, Leu347 and the aliphatic portion of Glu344. Previous mutational analysis highlighted the importance of Cet1 Pro245 and Trp247 side chains for function in vivo (Takase et al., 2000) and mutational studies on theC. albicans Cgt1 guanylyltransferase suggested that Trp309, Leu312 and Leu319 side chains contributed to Cgt1’s essential functions in vivo and to CaCet1 binding in vitro (Fabrega et al., 2003). Cgt1 Trp309, Leu312 and Leu319 are analogous to Ceg1 Trp337, Leu340 and Leu347. In the present study, the importance of contacts between Cet1 and Ceg1 observed in our structure were assessed for function in vivo by constructing mutantceg1 alleles and testing their ability to complement aΔceg1 yeast strain that expressed full-length Cet1 (CET1(1–549)) or aΔcet1Δceg1 yeast strain (Hausmann et al., 2001) that containedCET1(241–549), an allele that suffices for growth in vivo and is analogous to the Cet1 construct used in our crystal structure (Experimental Procedures). Consistent with our previous studies on Cgt1, strains containing W337H or L347Aceg1 alleles exhibited slow growth phenotypes at elevated temperatures inS. cerevisiae containing (CET1(1–549)) which became more severe in the strain containingCET1(241–549) (Figures 5C and 5D).

Previous studies on the Cgt1 guanylyltransferase highlighted the importance of Phe258 and Tyr278 side chains for function in vivo and for CaCet1 binding in vitro (Fabrega et al., 2003). The analogous residues in Ceg1 are Ile264 and Tyr284, both of which form one side of a complementary hydrophobic surface adjacent to Cet1 Pro245 and Ala248 (Figures 5A, 5B andFigure 6A). Consistent with interactions observed in our structure, I264A and Y284Aceg1 alleles exhibited slow growth phenotypes in theS. cerevisiae strain containingCET1(241–549) at elevated temperatures (Figure 5D). Two other tyrosine residues in Ceg1 (Tyr281 and Tyr282) were also selected for alanine substitution however these mutations did not result in growth defects in either strain. These results are consistent with our structure because Tyr281 and Try282 were not observed interacting with Cet1 and were presumed disordered within a loop between amino acids 265 and 284 (dashed line inFigures 5A and 5B). These results are also consistent with previous genetic and biochemical studies which suggested that individual alanine substitutions for analogous positions in Cgt1 (Tyr275 and Leu276) resulted in no detectable growth defects in vivo or for CaCet1 binding in vitro (Fabrega et al., 2003).

Previous mutational analysis within the conserved Cet1 WAQKW motif, namely K250A-W251A, resulted in loss of Cet1-Ceg1 interaction in vitro and elicited a temperature-sensitive growth phenotype in vivo that could be suppressed by over-expression of Ceg1 (Lehman et al., 1999). Cet1 amino acids Ala248, Gln249, Lys250 and Trp251 establish van der Waals and electrostatic contacts to Ceg1 side chain and main chain atoms from residues 345–350. In addition, this region of Cet1 contacts Ceg1 amino acid side chains in a helical-loop (HL) insertion that is not conserved with theC. albicans guanylyltransferase Cgt1 (Figure 6B). The remaining amino acid residues of the Cet1 N-terminal element make additional albeit less extensive contacts to Ceg1 residues Asp370, Asp371 and Leu373 (Figure 6A). The point mutations F308A, Q310A and F312A or a mutant allele that deleted the HL insertion (Δ304–323 or ΔHL) exhibited no growth defects in the strain expressing full length Cet1 (Figure 5C), however introduction of these alleles into the strain containingCET1(241–549) revealed that F312A did not suffice for growth as no colonies were obtained on media containing 0.75 mg/ml 5-fluoroorotic acid (5-FOA) after ten 10 days at 30 °C or 37 °C, and only small pinpoint colonies were obtained after 10 days at 23 °C (not shown). While strains containing F308A or Δ304–323 alleles were viable, both exhibited temperature sensitive growth defects at 23 °C or 37 °C (Figure 5D). These genetic data are consistent with contacts observed in our structure and with previous studies that support an essential role for interactions between the Cet1 WAQKW motif and the Ceg1 OB domain (Cho et al., 1998;Takase et al., 2000;Hausmann et al., 2001;Fabrega et al., 2003).

Production and purification of the Cet1-Ceg1 complexes

S. cerevisiae RNA triphosphatase,CET1(241–549) and RNA guanylyltransferaseCEG1 were amplified from genomic DNA obtained from a haploid W303-1A strain.CET1(241–549) was cloned into pET29b (Novagen), placing an ATG start codon at position 240. This plasmid was named pET29b-ScCet1(241–549). Full lengthS cerevisiae RNA guanylyltransferase,CEG1, was cloned into a pSMT3 TOPO vector (Mossessova and Lima, 2000; Invitrogen), prior to sub-cloning the His₆Smt3-ScCeg1 reading frame into pET15b to generate pET15b-Smt3ScCeg1. The vectors pET29b-ScCet1(241–549) and pET15b-Smt3ScCeg1 were co-transformed intoE. coli BL21 (DE3) CodonPlus RIL (Novagen). A 10 l culture was grown by fermentation at 37°C to an A₆₀₀ of 3 before 1 mM IPTG induction for 4 hours at 25°C. Cells were harvested by centrifugation, suspended in 50 mM Tris-HCl (pH 8.0) and 20% sucrose, 500 mM NaCl, 20 mM imidazole, 0.1% IGEPAL, 1 mM PMSF, 1 mM β-mercaptoethanol (BME), and 10 µg/ml DNAse, sonicated and insoluble material removed by centrifugation. His₆Smt3-Ceg1 was purified by metal-affinity chromatography (QIAGEN Ni-NTA Superflow resin) and Cet1 was co-purified via interaction with Ceg1. The His₆-Smt3 tag was removed by Ulp1 proteolysis on ice for 1 hour (Mossessova and Lima, 2000), and the complex was purified by gel filtration and anion exchange chromatography (Superdex200 and MonoQ, Pharmacia). Two distinct species containing both Cet1 and Ceg1 polypeptides were separated through both purification steps. Gel filtration profiles and SDS-PAGE analysis indicated that one was composed of two molecules of Cet1 and Ceg1, respectively, and the other was composed of two molecules of Cet1 and one molecule of Ceg1. Approximately 10 mg was obtained for each complex per liter ofE. coli fermentation culture as estimated by the Bradford assay (Bio-Rad). Both complexes were concentrated to 10 mg/ml in 50 mM NaCl, 10 mM Tris-HCl pH 8.0, 1 mM DTT. Aliquots were flash frozen in liquid nitrogen and stored at −80 °C.

To produce selenium-methionine labeled protein, pET29b-ScCet1(241–549) and pET15b-Smt3ScCeg1 were co-transformed intoE. coli B834 (Novagen) and grown in minimal media containing selenomethionine (Hendrickson et al., 1990). Cells grew at 37 °C until cell density reached an A₆₀₀ of ~2.0. Temperature was reduced to 30 °C and the culture was adjusted to 1 mM IPTG to induce protein expression. After 6 hours, cells were harvested and stored at −80 °C. The Cet1-Ceg1 protein obtained from these cultures was purified and crystallized as described for the native protein.

Crystallographic analysis

The Cet1-Ceg1 complex was crystallized by hanging drop vapor diffusion against a well solution that contained 1.0 M ammonium citrate, 0.1 M sodium citrate (pH 5.6), and 1.0% PEG (polyethylene glycerol) 4000 at 18°C. Large crystals (> 300 µM) were transferred to a solution containing well solution and 24% xylitol, incubated in this solution at 4 °C overnight, and flash cooled in liquid nitrogen. To obtain mercury derivatives, crystals were placed into solutions that contained crystallization media and 0.5 mM thimerosal at 4 °C overnight for 8 hours or for 16 hours. Crystals were cryo-protected and frozen as above. A complete data set was collected from native and mercury derivatized crystals at the Advanced Photon Source, SGX-CAT beamline (Argonne National Laboratory, Argonne, IL). Diffraction intensities were processed with DENZO and reduced using SCALEPACK (Otwinowski and Minor, 1997) and CCP4 (Collaborative Computational Project, 1994). SHARP was used to calculate phases to 3.2 Å (de La Fortelle and Bricogne, 1997). A model for the Cet1 dimer and its twin mate was generated by manually docking two Cet1 dimers into the electron density based on previous structures of Cet1 (Lima et al., 1999). The OB and NT domains fromC. albicans Cgt1 were used as a starting point for building the Ceg1 structure using the O program (Jones et al., 1991) to build missing segments in Ceg1 and Cet1.

A physical model for twinning emanates from analysis of the quaternary structure in the lattice as it is arranged as a trimer of Cet1-Ceg1 heterodimers which form a ring-like structure (Figure S3). The plane of the ring is perpendicular to the 6-fold axis and though the real space group is P6₃, the two-fold twinning operator (h+k, −k, −l) is perpendicular to the six-fold axis and makes the data appear consistent with P6₃22. Experimental phases determined in P6₃ revealed one orientation for each of the two Ceg1 molecules and two nearly orthogonal orientations for the Cet1 homodimer in the asymmetric unit (Figures S2 and S3). Because the twinning operator flips the pseudo-hexameric ring upside down, the structure and its twin mate align with perfect superposition of the respective Ceg1 molecules (thus the reason for observing density for Ceg1 in only one position) and two orientations for Cet1 that correspond to the two distinct positions observed in the experimental electron density for the Cet1 dimer (Figure S2). Although a 1:1 complex between Cet1 and Ceg1 is also possible in P6₃22, differences were observed for Cet1 amino acids 436–450 that break symmetry and suggest that the correct space group includes a heterotetramer in the asymmetric unit. Further corroborative evidence that P6₃ was the most probable space group was the observation that molecular replacement trials using Phaser generated clear rotational and translational solutions in P6₃22, but packing failed unless we allowed back bone clashes. On the other hand, molecular replacement in P6₃ generated clear solutions without steric consequences. The final scheme utilized tight NCS restraints for backbone atoms and loose NCS restraints for side chain atoms and by applying the twin operator (h+k, −k, −l) during refinement in Refmac (Murshudov et al., 1997). The final R_work and R_free values were 0.249 and 0.298, respectively. The model exhibits reasonable geometry for this resolution with 65.3% of residues in most favored, 30.6% of residues in additional allowed, 4.1% of residues in generously allowed, and no residues in disallowed regions of Ramachandran space, values that fall within accepted parameters at this resolution as determined by PROCHECK (Laskowski et al., 1993). Refinement required manual adjustments in the weighting scheme for geometry and x-ray terms resulting in slightly higher than expected r.m.s. deviations in bond length (0.017 Å), however random inspection of several recently determined high resolution structures (1.1–1.25 Å; PDB codes 3A9J, 3HS4, 3KOM, 3KHF, 3FBW, and 3HZA) refined using Refmac, Phenix and Shelx revealed values for r.m.s. deviation in bond lengths between 0.010–0.017 Å. A composite simulated annealing omit map is shown covering several regions of our model (Figure S4).

As an alternative to twinning, we attempted to refine the Cet1-Ceg1 model assuming that both Cet1 homodimers were superimposed because of static disorder in the lattice. This was initiated by modeling two Ceg1 chains at full occupancy and the two conformations of the Cet1 homodimer by four chains (A and B, A’ and B’) with each chain at half occupancy. Because A and B’ as well as B and A’ overlap (Figures S2A, S2B, and S3C), refinement of the six chains was enabled by turning off repulsive forces between the overlapping Cet1 molecules in CNS (Brunger et al., 1998). The structure was refined using NCS restraints and harmonic backbone restraints for regions of helical or beta secondary structure at 3.0 Å using the twinned data to a R_work and R_free of 0.275 and 0.337, respectively, however this model, and the statistical analysis which indicated that the data were perfectly twinned, failed to provide a physical explanation for the two overlapping Cet1 protomers.

Mutagenesis and yeast growth assays

The plasmid pGYCE-358 contained theCEG1 reading frame under the control of theCEG1 endogenous promoter (Schwer and Shuman, 1994). Mutations to replace solvent exposed hydrophobic residues in the Ceg1 OB-fold domain and residues in contact with Cet1 were generated by PCR. A deletion mutant was generated for Ceg1 that removed a helix and loop (Δ304–323) observed in interactions with Cet1. Plasmids containingCEG1 mutations were pGYCE-358-I264A, pGYCE-358-Y281A, pGYCE-358-Y282A, pGYCE-358-Y284A, pGYCE-358-F308A, pGYCE-358-Q310A, pGYCE-358-F312A, pGYCE-358-W337H, and pGYCE-358-L347A. The plasmid encoding the internal deletion inCEG1 was pGYCE-358-Δ304–323. The effects of mutations on cell growth were tested by transforming respective pGYCE-358 plasmids into theS. cerevisiae strain YBS2 (MATa, leu2, lys2, trp1, ceg1::hisG, pGYCE-360) whereceg1Δ is complemented by theCEN URA3 shuffling plasmid pGYCE-360 that contained wild-typeCEG1 under control of its endogenous promoter.

The plasmid pRS412-CET1(241–549) was constructed by amplifying theCET1(241–549) fragment by PCR from genomic DNA of theS. cerevisiae strain W303-1A. TheCET1(241–549) fragment was flanked by the endogenous 5’ (520 bp) and 3’ (530 bp) UTR elements located adjacent to theCET1 coding region. The effects for respective mutant alleles on cell growth were tested by co-transforming respective pGYCE-358 and pRS412-CET1(241–539) plasmids into theS. cerevisiae strain YBS50 (MATa, leu2, ade2, trp1, his3, ura3, can1 ceg1::hisG, cet1::LEU2, p360-CET1/CEG1) in whichcet1Δceg1Δ was complemented by theCEN URA3 plasmid p360-CET1/CEG1 that contained wild-typeCET1 andCEG1 (Hausmann et al., 2001).

Strains were grown on agar plates containing synthetic medium lacking either tryptophan or tryptophan and adenine to select colonies that contained the respective plasmids. Individual colonies were streaked on agar plates that contained 0.75 mg/ml 5-fluoroorotic acid (5-FOA) to select for loss of the respective URA3 plasmids. Mutations deemed insufficient for supporting cell growth were those that failed to yield 5–FOA resistant colonies after incubation for 10 days at 23°C, 30°C or 37°C. Viable strains were grown in YPAD broth to an A₆₀₀ of 0.6 to 0.8, adjusted to an A600 of 0.5 in 15% glycerol and stored at −80 °C. Each strain and the respective serial 10-fold dilutions (10⁻¹, 10⁻² and 10⁻³) were spotted on YPAD agar plates. All the stains were scored after growth at 23°C, 30°C and 37°C for 3 to 4 days.

1. Benarroch D, Smith P, Shuman S. Characterization of a trifunctional mimivirus mRNA capping enzyme and crystal structure of the triphosphatase domain. Structure. 2008;16:501–512. doi: 10.1016/j.str.2008.01.009. [DOI] [PubMed] [Google Scholar]
2. Brunger AT, Adams PD, Clore GM, DeLano WL, Gros P, Grosse-Kunstleve RW, Jiang JS, Kuszewski J, Nilges M, Pannu NS, Read RJ, Rice LM, Simonson T, Warren GL. Crystallography & NMR system: A new software suite for macromolecular structure determination. Acta. Crystallogr. D Biol. Crystallogr. 1998;54:905–921. doi: 10.1107/s0907444998003254. [DOI] [PubMed] [Google Scholar]
3. Changela A, Ho CK, Martins A, Shuman S, Mondragon A. Structure and mechanism of the RNA triphosphatase component of mammalian mRNA capping enzyme. EMBO J. 2001;20:2575–2586. doi: 10.1093/emboj/20.10.2575. [DOI] [PMC free article] [PubMed] [Google Scholar]
4. Chiu YL, Ho CK, Saha N, Schwer B, Shuman S, Rana TM. Tat stimulates cotranscriptional capping of HIV mRNA. Mol. Cell. 2002;10:585–597. doi: 10.1016/s1097-2765(02)00630-5. [DOI] [PubMed] [Google Scholar]
5. Cho EJ, Rodriguez CR, Takagi T, Buratowski S. Allosteric interactions between capping enzyme subunits and the RNA polymerase II carboxy-terminal domain. Genes Dev. 1998;12:3482–3487. doi: 10.1101/gad.12.22.3482. [DOI] [PMC free article] [PubMed] [Google Scholar]
6. Cho EJ, Takagi T, Moore CR, Buratowski S. mRNA capping enzyme is recruited to the transcription complex by phosphorylation of the RNA polymerase II carboxy-terminal domain. Genes Dev. 1997;11:3319–3326. doi: 10.1101/gad.11.24.3319. [DOI] [PMC free article] [PubMed] [Google Scholar]
7. Corden JL. Tails of RNA polymerase II. Trends Biochem. Sci. 1990;15:383–387. doi: 10.1016/0968-0004(90)90236-5. [DOI] [PubMed] [Google Scholar]
8. de La Fortelle E, Bricogne G. Maximum-likelihood heavy-atom parameter refinement for multiple isomorphous replacement and multiwavelength anomalous diffraction methods. Methods Enzymol. 1997;276:472–494. doi: 10.1016/S0076-6879(97)76073-7. [DOI] [PubMed] [Google Scholar]
9. Dahmus ME. The role of multisite phosphorylation in the regulation of RNA polymerase II activity. Prog. Nucleic Acid Res. Mol. Biol. 1994;48:143–179. doi: 10.1016/s0079-6603(08)60855-7. [DOI] [PubMed] [Google Scholar]
10. Egloff S, Murphy S. Cracking the RNA polymerase II CTD code. Trends Genet. 2008;24:280–288. doi: 10.1016/j.tig.2008.03.008. [DOI] [PubMed] [Google Scholar]
11. Fabrega C, Hausmann S, Shen V, Shuman S, Lima CD. Structure and mechanism of mRNA cap (guanine-N7) methyltransferase. Mol. Cell. 2004;13:77–89. doi: 10.1016/s1097-2765(03)00522-7. [DOI] [PubMed] [Google Scholar]
12. Fabrega C, Shen V, Shuman S, Lima CD. Structure of an mRNA capping enzyme bound to the phosphorylated carboxy-terminal domain of RNA polymerase II. Mol. Cell. 2003;11:1549–1561. doi: 10.1016/s1097-2765(03)00187-4. [DOI] [PubMed] [Google Scholar]
13. Furuichi Y, Shatkin AJ. Viral and cellular mRNA capping: past and prospects. Adv. Virus Res. 2000;55:135–184. doi: 10.1016/S0065-3527(00)55003-9. [DOI] [PMC free article] [PubMed] [Google Scholar]
14. Gu M, Lima CD. Processing the message: structural insights into capping and decapping mRNA. Curr. Opin. Struct. Biol. 2005;15:99–106. doi: 10.1016/j.sbi.2005.01.009. [DOI] [PubMed] [Google Scholar]
15. Hakansson K, Doherty AJ, Shuman S, Wigley DB. X-ray crystallography reveals a large conformational change during guanyl transfer by mRNA capping enzymes. Cell. 1997;89:545–553. doi: 10.1016/s0092-8674(00)80236-6. [DOI] [PubMed] [Google Scholar]
16. Hakansson K, Wigley DB. Structure of a complex between a cap analogue and mRNA guanylyl transferase demonstrates the structural chemistry of RNA capping. Proc. Natl. Acad. Sci. U.S.A. 1998;95:1505–1510. doi: 10.1073/pnas.95.4.1505. [DOI] [PMC free article] [PubMed] [Google Scholar]
17. Hausmann S, Ho CK, Schwer B, Shuman S. An essential function of Saccharomyces cerevisiae RNA triphosphatase Cet1 is to stabilize RNA guanylyltransferase Ceg1 against thermal inactivation. J. Biol. Chem. 2001;276:36116–36124. doi: 10.1074/jbc.M105856200. [DOI] [PubMed] [Google Scholar]
18. Hendrickson WA, Horton JR, LeMaster DM. Selenomethionyl proteins produced for analysis by multiwavelength anomalous diffraction (MAD): a vehicle for direct determination of three-dimensional structure. EMBO J. 1990;9:1665–1672. doi: 10.1002/j.1460-2075.1990.tb08287.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
19. Hayward S, Berendsen HJ. Systematic analysis of domain motions in proteins from conformational change: new results on citrate synthase and T4 lysozyme. Proteins. 1998;30:144–154. [PubMed] [Google Scholar]
20. Ho CK, Schwer B, Shuman S. Genetic, physical, and functional interactions between the triphosphatase and guanylyltransferase components of the yeast mRNA capping apparatus. Mol. Cell. Biol. 1998;18:5189–5198. doi: 10.1128/mcb.18.9.5189. (a) [DOI] [PMC free article] [PubMed] [Google Scholar]
21. Ho CK, Sriskanda V, McCracken S, Bentley D, Schwer B, Shuman S. The guanylyltransferase domain of mammalian mRNA capping enzyme binds to the phosphorylated carboxyl-terminal domain of RNA polymerase II. J. Biol. Chem. 1998;273:9577–9585. doi: 10.1074/jbc.273.16.9577. (b) [DOI] [PubMed] [Google Scholar]
22. Ho CK, Lehman K, Shuman S. An essential surface motif (WAQKW) of yeast RNA triphosphatase mediates formation of the mRNA capping enzyme complex with RNA guanylyltransferase. Nucleic Acids Res. 1999;27:4671–4678. doi: 10.1093/nar/27.24.4671. [DOI] [PMC free article] [PubMed] [Google Scholar]
23. Ho CK, Shuman S. Distinct roles for CTD Ser-2 and Ser-5 phosphorylation in the recruitment and allosteric activation of mammalian mRNA capping enzyme. Mol Cell. 1999;3:405–411. doi: 10.1016/s1097-2765(00)80468-2. [DOI] [PubMed] [Google Scholar]
24. Jones TA, Zou JY, Cowan SW, Kjeldgaard Improved methods for building protein models in electron density maps and the location of errors in these models. Acta. Crystallogr. A. 1991;47:110–119. doi: 10.1107/s0108767390010224. [DOI] [PubMed] [Google Scholar]
25. Jove R, Manley JL. Transcription initiation by RNA polymerase II is inhibited by S-adenosylhomocysteine. Proc. Natl. Acad. Sci. U. S. A. 1982;79:5842–5846. doi: 10.1073/pnas.79.19.5842. [DOI] [PMC free article] [PubMed] [Google Scholar]
26. Komarnitsky P, Cho EJ, Buratowski S. Different phosphorylated forms of RNA polymerase II and associated mRNA processing factors during transcription. Genes Dev. 2000;14:2452–2460. doi: 10.1101/gad.824700. [DOI] [PMC free article] [PubMed] [Google Scholar]
27. Laskowski RA, MacArthur MW, Moss DS, Thornton JM. PROCHECK: a program to check the stereochemical quality of protein structures. J. App. Cryst. 1993;26:283–291. [Google Scholar]
28. Lehman K, Schwer B, Ho CK, Rouzankina I, Shuman S. A conserved domain of yeast RNA triphosphatase flanking the catalytic core regulates self-association and interaction with the guanylyltransferase component of the mRNA capping apparatus. J. Biol. Chem. 1999;274:22668–22678. doi: 10.1074/jbc.274.32.22668. [DOI] [PubMed] [Google Scholar]
29. Lima CD, Wang LK, Shuman S. Structure and mechanism of yeast RNA triphosphatase: an essential component of the mRNA capping apparatus. Cell. 1999;99:533–543. doi: 10.1016/s0092-8674(00)81541-x. [DOI] [PubMed] [Google Scholar]
30. McCracken S, Fong N, Rosonina E, Yankulov K, Brothers G, Siderovski D, Hessel A, Foster S, Shuman S, Bentley DL. 5'-Capping enzymes are targeted to pre-mRNA by binding to the phosphorylated carboxy-terminal domain of RNA polymerase II. Genes Dev. 1997;11:3306–3318. doi: 10.1101/gad.11.24.3306. [DOI] [PMC free article] [PubMed] [Google Scholar]
31. Mossessova E, Lima CD. Ulp1-SUMO crystal structure and genetic analysis reveal conserved interactions and a regulatory element essential for cell growth in yeast. Mol. Cell. 2000;5:865–876. doi: 10.1016/s1097-2765(00)80326-3. [DOI] [PubMed] [Google Scholar]
32. Murshudov GN, Vagin AA, Dodson EJ. Refinement of Macromolecular Structures by the Maximum-Likelihood method. Acta Cryst. 1997;D53:240–255. doi: 10.1107/S0907444996012255. [DOI] [PubMed] [Google Scholar]
33. Otwinowski Z, Minor W. Processing of X-ray diffraction data collected in oscillation mode. Methods Enzymol. 1997;276:307–326. doi: 10.1016/S0076-6879(97)76066-X. [DOI] [PubMed] [Google Scholar]
34. Padilla J, Yeates TO. A statistic for local intensity differences: robustness to anisotropy and pseudo-centering and utility for detecting twinning. Acta Crystallogr. 2003;D59:1124–1130. doi: 10.1107/s0907444903007947. [DOI] [PubMed] [Google Scholar]
35. Pei Y, Hausmann S, Ho CK, Schwer B, Shuman S. The length, phosphorylation state, and primary structure of the RNA polymerase II carboxyl-terminal domain dictate interactions with mRNA capping enzymes. J. Biol. Chem. 2001;276:28075–28082. doi: 10.1074/jbc.M102170200. [DOI] [PubMed] [Google Scholar]
36. Phatnani HP, Greenleaf AL. Phosphorylation and functions of the RNA polymerase II CTD. Genes Dev. 2006;20:2922–2936. doi: 10.1101/gad.1477006. [DOI] [PubMed] [Google Scholar]
37. Rasmussen EB, Lis JT. In vivo transcriptional pausing and cap formation on three Drosophila heat shock genes. Proc. Natl. Acad. Sci. U. S. A. 1993;90:7923–7927. doi: 10.1073/pnas.90.17.7923. [DOI] [PMC free article] [PubMed] [Google Scholar]
38. Schroeder SC, Schwer B, Shuman S, Bentley D. Dynamic association of capping enzymes with transcribing RNA polymerase II. Genes Dev. 2000;14:2435–2440. doi: 10.1101/gad.836300. [DOI] [PMC free article] [PubMed] [Google Scholar]
39. Schroeder SC, Zorio DA, Schwer B, Shuman S, Bentley D. A function of yeast mRNA cap methyltransferase, Abd1, in transcription by RNA polymerase II. Mol. Cell. 2004;13:377–387. doi: 10.1016/s1097-2765(04)00007-3. [DOI] [PubMed] [Google Scholar]
40. Schwer B, Shuman S. Mutational analysis of yeast mRNA capping enzyme. Proc. Natl. Acad. Sci. U. S. A. 1994;91:4328–4332. doi: 10.1073/pnas.91.10.4328. [DOI] [PMC free article] [PubMed] [Google Scholar]
41. Shuman S. The mRNA capping apparatus as drug target and guide to eukaryotic phylogeny. Cold Spring Harb. Symp. Quant. Biol. 2001;66:301–312. doi: 10.1101/sqb.2001.66.301. [DOI] [PubMed] [Google Scholar]
42. Shuman S, Lima CD. The polynucleotide ligase and RNA capping enzyme superfamily of covalent nucleotidyltransferases. Curr. Opin. Struct. Biol. 2004;14:757–764. doi: 10.1016/j.sbi.2004.10.006. [DOI] [PubMed] [Google Scholar]
43. Takase Y, Takagi T, Komarnitsky PB, Buratowski S. The essential interaction between yeast mRNA capping enzyme subunits is not required for triphosphatase function in vivo. Mol Cell Biol. 2000;20:9307–9316. doi: 10.1128/mcb.20.24.9307-9316.2000. [DOI] [PMC free article] [PubMed] [Google Scholar]
44. Takagi T, Cho EJ, Janoo RT, Polodny V, Takase Y, Keogh MC, Woo SA, Fresco-Cohen LD, Hoffman CS, Buratowski S. Divergent subunit interactions among fungal mRNA 5'-capping machineries. Eukaryotic Cell. 2002;1:448–457. doi: 10.1128/EC.1.3.448-457.2002. [DOI] [PMC free article] [PubMed] [Google Scholar]
45. Takagi T, Moore CR, Diehn F, Buratowski S. An RNA 5'-triphosphatase related to the protein tyrosine phosphatases. Cell. 1997;89:867–873. doi: 10.1016/s0092-8674(00)80272-x. [DOI] [PubMed] [Google Scholar]
46. Wen Y, Yue Z, Shatkin AJ. Mammalian capping enzyme binds RNA and uses protein tyrosine phosphatase mechanism. Proc. Natl. Acad. Sci. U.S.A. 1998;95:12226–12231. doi: 10.1073/pnas.95.21.12226. [DOI] [PMC free article] [PubMed] [Google Scholar]
47. Yue Z, Maldonado E, Pillutla R, Cho H, Reinberg D, Shatkin AJ. Mammalian capping enzyme complements mutant Saccharomyces cerevisiae lacking mRNA guanylyltransferase and selectively binds the elongating form of RNA polymerase II. Proc. Natl. Acad. Sci. U. S. A. 1997;25:12898–12903. doi: 10.1073/pnas.94.24.12898. [DOI] [PMC free article] [PubMed] [Google Scholar]

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