REGULATION OF U1 snRNP / 5’ SPLICE SITE INTERACTIONS DURING PRE-mRNA SPLICING IN SACCHAROMYCES CEREVISIAE
DISSERTATION
Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University
By
Leah R. Stands
* * * * *
The Ohio State University 2003
Dissertation Committee:
Dr. Tien-Hsien Chang, Adviser Approved by Dr. Charles Daniels
Dr. Paul Herman Adviser Dr. Lee F. Johnson Department of Molecular Genetics
ABSTRACT
Intron removal is a complex process carried out by a dynamic cellular machine termed the spliceosome. To form a functional spliceosome, five small nuclear ribonucleoprotein particles (snRNPs) must sequentially bind to a precursor mRNA (pre-mRNA) and then be rearranged via an ordered series of
RNA-RNA interactions. Many of these RNA-RNA interactions are mutually exclusive, i.e. the formation of one requires the disruption of another. For example, the 5’ splice site (SS) of the pre-mRNA is initially recognized by base pairing with the U1 snRNA. This interaction must be disrupted to allow the U6 snRNA to base pair with the 5’ SS. Such RNA-RNA rearrangements are likely to require factors that facilitate base pairing as well as others that disrupt the interactions. The disruption of these interactions is likely catalyzed by members of the DExD/H box family of putative RNA helicases, enzymes that catalyze the
ii disruption of RNA duplexes and are involved in many cellular processes including pre-mRNA splicing.
Prp28p is a putative RNA helicase required during the early stages of pre- mRNA splicing. Specifically, Prp28p is required for the switch of U1 for U6 at the
5’ SS and may promote this switch by destabilize the U1/5’ SS RNA duplex.
Interestingly, mutant versions of the U1 snRNP-specific U1-C protein can eliminate the requirement for the normally essential Prp28p. The mutant U1-C proteins may weaken the U1/5’ SS interaction, thus eliminating the need for
Prp28p to destabilize the RNA duplex. Therefore Prp28p may serve not only to disrupt an RNA duplex, but to also counteract the stabilizing effect of the U1-C protein.
Presented here are studies aimed at understanding the complex target of
Prp28p and its function in splicing. In addition to continuing analysis of alterations in the U1-C protein that allow bypass of Prp28p, I have identified a specific domain within the U5 snRNP protein Prp8p that may assist Prp28p in displacing U1 from the 5’ SS. Alterations in two additional U1 snRNP proteins,
Prp42p and Snu71p, were also found to bypass Prp28p, indicating that these proteins may also be targeted by this putative helicase. Alterations in several Sm
iii proteins known to disrupt the U1/5’ SS association cannot bypass Prp28p, indicating that the bypass abilities of U1-C, Prp42p and Snu71p are specific and that these proteins likely represent true targets of Prp28p. I present a model in which the proteins U1-C, Prp42p, and Snu71p work to facilitate the U1/5’ SS interaction while Prp28p and Prp8p work antagonistically to disrupt this association. My data also contributes to a growing body of evidence signaling a shift in the way RNA helicase function is viewed. These enzymes may not target only RNA duplexes, but instead may modify proteins and RNA, functioning as
RNPases to alter both protein and RNA contacts within RNP complexes.
iv Dedicated to my family
v ACKNOWLEDGMENTS
I wish to thank my adviser, Dr. Chang, for his guidance during my graduate training as well as the members of the Chang lab for their support and assistance. I also thank Dr. Charles Daniels, Dr. Paul Herman, and Dr. Lee
Johnson for their assistance as my committee members as well as Dr. David
Brow, Dr. Cathy Collins, Dr. Hansen Du, Dr. Christine Guthrie, Dr. Magda
Konarska, Dr. Andreas Kuhn, Dr. Michael Rosbash and Dr. Jon Staley for sharing data and reagents. Finally, I must thank my family and friends, especially my classmates Vicki, Anil and Linda, for their support and encouragement during my graduate education.
vi VITA
February 28, 1975………………………Born – Berea, Ohio
1997………………………………………B.S. Biology, Ohio Northern University
1997 – present…………………………..Graduate Teaching and Research Associate, Department of Molecular Genetics, The Ohio State University
PUBLICATIONS
Chen, J. Y.-F., Stands, L., Staley, J.P., Jackups, R.R., Latus, L.J., and Chang, T.-H. 2001. Specific alterations of U1-C protein or U1 small nuclear RNA can eliminate the requirement of Prp28p, an essential DEAD box splicing factor. Mol. Cell 7:227-232.
FIELDS OF STUDY
Major Field: Molecular Genetics
vii TABLE OF CONTENTS
Page
Abstract………………………………………………………………………………..….ii
Dedication……………………………………………………………………………..…v
Acknowledgments………………………………………………………………………vi
Vita…………………………………………………………………………….…………vii
List of Tables…………………………………………………………………………...xii
List of Figures………………………………………………………………………….xiii
Chapters:
1. Introduction………………………………………………………………………….1
1.1 Pre-mRNA Splicing………………………………………………….…....1 1.2 The Chemistry of pre-mRNA Splicing……………………………….…..3 1.3 The Splicing Machinery……………………………………………………4 1.3.1 The Spliceosome………………………………………………...4 1.3.2 snRNAs and snRNPs……………………………………………5 1.3.2.1 The U1 snRNP…………………………………………8 1.3.2.2 The U1-C Protein……………………………………..10 1.4 The Dynamic Nature of the Spliceosome…………………………...….13 1.4.1 Assembly of the Spliceosome……………………………...….13 1.4.2 DExD/H box Proteins………………………………………...…15 1.4.2.1 DExD/H Box Proteins in splicing…………………….23
viii 1.4.3 The Holospliceosome Hypothesis…………………………….27 1.4.4 Prp28p…………………………………………………………...29 1.4.5 Recognition of the 5’ Splice Site………………………………33 1.4.6 Prp8p and the Catalytic Core of the Spliceosome….……….38 1.5 Goal of This Work…………………………………………………………46
2. Materials and Methods
2.1 Primer Extension…………………………………………………………..63 2.2 In vitro Splicing…………………………………………………………….64
2.3 Purification of His6-Prp28p and Production of anti-Prp28p Antibody..65 2.4 Testing U1 snRNA Mutations for Bypass of PRP28…………………..67 2.5 Testing the Growth Phenotypes of U1-C L-13 Variants………………69 2.6 RT-PCR Confirmation of Splicing Defects Identified by Microarray Analysis of prp28∆ YHC1-1 Cells………………………….70 2.7 Screening for Mutations Synthetically Lethal with prp28∆ YHC1-1….73 2.8 Characterization and Cloning of prp8-501……………………………...74 2.9 Testing PRP8 Mutations for Synthetic Lethality with prp28∆ YHC1-1……………………………………………………………76 2.10 Back-crossing of Bypass Suppressor Strains………………………...78 2.11 Identification of prp42-101 and snu71-101 as a Bypass Suppressors of PRP28………………………………………………….79 2.12 Cloning of Wild-type and Mutant PRP42……………………………...80 2.13 Cloning of Wild-type and Mutant SNU71…………………………...…82 2.14 Analyzing the Growth Phenotype of prp28∆ prp42-101……………..83 2.15 prp42-101 Does not Produce a Growth Defect on its Own………….83 2.16 Deletion of PRP42 Cannot Bypass PRP28…………………………...84 2.17 prp42-101 and snu71-101 are Synthetically Lethal with YHC1 Bypass Alleles…………………………………………………….85 2.18 Alleles of prp8 are Synthetically Lethal with prp42-101…………..…89
2.19 Testing Tail-truncated Sm Proteins for bypass of Prp28p…………..90 2.20 Deletion of MUD1 Cannot Bypass PRP28……………………………91
ix 3. Results
3.1 Investigation of Alterations in the U1-C Protein and the U1 snRNA that Allow Bypass ofPRP28……………………………….106 3.1.1 Determination of in vivo and in vitro Splicing Defects in the prp28∆ YHC1-1 Strain………………………………...107
3.1.2 Recombinant His6-Prp28p Relieves Splicing Defects in vitro ………………………………………………………….109 3.1.3 Alterations in the U1 snRNA Bypass PRP28……………….110 3.1.4 Investigating the Growth Phenotypes of the U1-C L13 Variants……………………………………………………112 3.1.5 Verification of Splicing Defects in Transcripts that are Highly Sensitive to prp28∆ YHC1-1……………………113 3.2 Prp8p Collaborates with Prp28p During the U1/5’ SS Rearrangements…………………………………………………………115 3.2.1 prp8-501 is Synthetically Lethal with prp28∆ YHC1-1…….116 3.2.2 Mutations Throughout PRP8 are Synthetically Lethal with prp28∆ YHC1-1…………………………………………..118 3.3 Identification and Preliminary Characterization of prp42-101 and snu71-101 as Bypass Suppressors of PRP28………………..…119 3.3.1 Novel Alleles of PRP42 and SNU71 Bypass the Requirement for Prp28p………………………………….125 3.3.2 The prp28∆ prp42-101 Strain Displays a Cold-sensitive Growth Phenotype………………………..…127 3.3.3 prp42-101 Does not Cause a Growth Defect on its Own…128 3.3.4 Deletion of prp42 Cannot Bypass PRP28………………….129 3.3.5 Analysis of the in vitro Splicing Defect in prp28∆ prp42-101……………………………………………..130 3.3.6 Synthetic Lethality of prp42-101 and snu71-101 with YHC1 Bypass Alleles……………………………………131 3.3.7 Alleles of prp8 are Synthetically Lethal with prp28∆ prp42-101……………………………………………..132
x 3.4 Bypass of PRP28 is a Specific Event………………………………….133 3.4.1 Truncation of SmB, SmD1 and SmD3 C-terminal Tails Cannot Bypass Prp28p…………………………………136 3.4.2 Deletion of MUD1 Cannot Bypass PRP28………………….137
4. Discussion
4.1 Investigation of Alterations in the U1-C Protein and the U1 snRNA that Allow Bypass of PRP28………………………………177 4.1.1 Characterization of the YHC1-1 Bypass Suppressor……...177 4.1.2 Alterations in the U1 snRNA Bypass PRP28………………178 4.1.3 Growth Phenotypes of the U1-C L13 Variants…………….180 4.1.4 A Novel Role for U1-C at the 5’ SS…………………………183 4.2 Novel Mutations in PRP42 and SNU71 Can Bypass PRP28……….185 4.2.1 prp42-101 and snu71-101 May Destabilize the U1/5’ SS Duplex……………………………………………….185 4.2.2 prp42-101 is Synthetically Lethal with Alleles of PRP8……186 4.2.3 Protein Interaction Domains May be Affected by prp42-101 and snu71-101………………………………….…188 4.2.4 Prp42p May be Required for the Efficient Splicing of a Subset of Transcripts………………………………….…189 4.3 Bypass of PRP28 is a Specific Event………………………………….195 4.3.1 Tail-truncated Versions of Three Sm Proteins Cannot Bypass PRP28………………………………………..195 4.3.2 Deletion of MUD1 Cannot Bypass PRP28………………….197
4.4 The Function of Prp8p at the Catalytic Core………………………….199 4.4.1 A Specific Domain Within Prp8p Appears to Aid in the Removal of U1 From the 5’ SS………………………….199 4.4.2 A Role for Prp8p in Spliceosomal Activation…………….…202 4.5 Prospectus………………………………………………………………..209
Bibliography…………………………………………………………………………..215
xi LIST OF TABLES
Table Page
2.1 Yeast Strains…………………………………………………………………..92
2.2 Plasmids………………………………………………………………………..96
3.1 PRP8 Mutations Synthetically Lethal with prp28∆ YHC1-1 and prp28∆ prp42-101……………………………………………………….158
xii LIST OF FIGURES
Figure Page 1.1 Selected types of alternative splicing…………………………………………..48 1.2 Consensus sequences of splicing signals……………………………………..50 1.3 The chemical mechanism of pre-mRNA splicing……………………………...51 1.4 Secondary structures of the human and yeast U1 snRNAs………………....52 1.5 The spliceosome cycle…………………………………………………………..54 1.6 Conserved sequence motifs of DExD/H box proteins………………………..56 1.7 Summary of known or suggested functions of DExD/H box protein motifs………………………………………………………………….…..58 1.8 DExD/H box proteins in pre-mRNA splicing…………………………………...59 1.9 Map of proposed functional domains within Prp8p…………………………...61 3.1 Primer extension analysis of splicing in prp28∆ YHC1-1 cells……………..138 3.2 In vitro splicing in prp28∆ YHC1-1 extracts…………………………………..139 3.3 Time course study of splicing at 25˚C in wild-type and prp28∆ YHC1-1 extracts………………………………………………………..141
3.4 Purification of recombinant His6-Prp28p and analysis of antibody production……………………………………………………………..143
3.5 Recombinant His6-Prp28p rescues splicing defects in vitro………………..144 3.6 Mutations in the U1 snRNA can bypass PRP28…………………………….145 3.7 Model for bypass of Prp28p by alterations in U1-C…………………………146 3.8 Growth phenotypes of the YHC1 L-13 bypass mutants in a yhc1∆ strain..147 3.9 Growth phenotypes of the YHC1 L-13 bypass mutants during bypass…...149 3.10 Agarose gel analysis of RT-PCR products…………………………………..151 3.11 Diagram of screen to identify mutations synthetically lethal with prp28∆ YHC1-1…………………………………………………………………152 3.12 Location of altered amino acids due to prp8-501 mutations……………….154 3.13 Mutations altering a distinct region of Prp8p are synthetically lethal with prp28∆ YHC1-1………………………………………………….…156 3.14 Map of Prp42p………………………………………………………………….161
xiii 3.15 Map of Snu71p…………………………………………………………..……..163 3.16 Growth phenotype of strains containing prp42-101………………………...165 3.17 Time course study of splicing in wild-type and prp28∆ prp42-101 extracts…………………………………………………….167 3.18 prp42-101 is synthetically lethal with bypass alleles of YHC1…………….169 3.19 snu71-101 is synthetically lethal with bypass alleles of YHC1…………….171 3.20 Mutations altering a distinct region of Prp8p are synthetically lethal with prp28∆ prp42-101………………………………………………….173 3.21 C-terminal truncations of SM proteins B, D1 and D3 cannot bypass Prp28p………………………………………………………………….175 3.22 Deletion of MUD1 cannot bypass Prp28p…………………………………...176
xiv CHAPTER 1
INTRODUCTION
1.1 Pre-mRNA Splicing
The majority of genes in most eukaryotes are discontinuous, meaning that their coding sequences are interrupted by non-coding sequences called introns.
For proper gene expression to take place, the non-coding sequences must be removed and the coding sequences properly joined. Many transcripts contain multiple introns that can undergo alternative splicing in a multitude of patterns to allow different forms of the same gene to be expressed, often in a temporally controlled and tissue-specific manner (Horowitz and Krainer, 1994; Maniatis,
1991) (figure 1.1). Intron removal occurs in a process called nuclear pre- messenger RNA (pre-mRNA) splicing after the gene has been transcribed.
Conserved sequence elements within the primary RNA transcript provide the signals required for precise intron removal. These sequences include 5’ and 3’
1 splice sites (SS), a branch point sequence containing a strictly conserved adenosine residue, and a stretch of pyrimidines that is located between the branch point and the 3’ splice site (figure 1.2).
Interestingly, although the conserved sequence at the 5’ and 3’ splice sites and branch point are similar in all organisms, these signals are much more conserved in the yeast Saccharomyces cerevisiae than in mammals. The 5’ SS sequence signal in yeast is almost always GUAUGU while in humans only the first two positions (i.e., GU) are very highly conserved. Similarly, the branch point sequence found in yeast introns is almost always UACUAAC, but the sequence is very degenerate in mammalian introns. In both yeast and mammals, the 3’ SS signal is rather short, consisting of a pyrimidine (U or C) followed by AG. The differences in signal sequence conservation likely reflect the relative complexity of the systems. As a simple, single-celled eukaryote, yeast have few introns and no alternative splicing. Their introns can therefore have very strong, clear-cut signals. Higher organisms that utilize alternative splicing, however, have a number of weaker signals that allow a choice between their usage under various conditions.
2 1.2 The Chemistry of pre-mRNA Splicing
Splicing occurs by a two step trans-esterification mechanism (figure 1.3)
(Ruby and Abelson, 1991; Sharp et al., 1987). In the first step, the phosphodiester bond at the 5’ SS is cleaved by a nucleophilic attack by the 2’ hydroxyl group of the conserved adenosine (A) located at the intron branch point.
This generates a 2’- 5’ phosphodiester bond between the branch site and the 5’ end of the intron as well as a free 3’ hydroxyl group on the 5’ exon. The cleavage also results in two RNA molecules, including a free exon 1 and a “lariat” intron-exon 2. In the second step, the free 3’ hydroxyl on the end of the 5’ exon attacks the phosphodiester bond at the 3’ splice site, resulting in the joining of the exons and release of the lariat intron. The basic chemistry of pre-mRNA splicing is similar to that of the group II introns, also called self-splicing introns, that are found in organelles. This group of introns catalyze their own removal and require no protein cofactors (Cech, 1990; Michel and Jacquier, 1987). In contrast to these self-splicing introns, nuclear pre-mRNA splicing requires an input of energy and protein cofactors that are organized into a large ribonucleoprotein particle called the spliceosome.
3 The study of pre-mRNA splicing has relied on a combination of genetic and biochemical approaches. Detection of genetic interactions among splicing factors has proven valuable in identifying proteins and RNAs that have critical functions in the splicing pathway. Mutational analysis of transcripts and snRNAs also provides invaluable information about molecular interactions during splicing.
The development of in vitro splicing assays using reporter transcripts and carefully purified cell extracts provided a critical tool to investigate the mechanisms of splicing in both yeast and mammalian systems (Hernandez and
Keller, 1983; Krainer et al., 1984; Lin et al., 1985).
1.3 The Splicing Machinery
1.3.1 The Spliceosome
Splicing is accomplished in a complex cellular machine called the spliceosome, which orchestrates the removal of introns from pre-mRNAs. Since the discovery that a large complex was responsible for splicing, the list of spliceosome components has grown to include five small nuclear RNAs
(snRNAs) and over 75 proteins ( Brody and Abelson, 1985; Brow, 2002). Each of the five snRNAs (U1, U2, U4, U5 and U6) is found in a complex with a number
4 of proteins to form small nuclear ribonucleoprotein particles or snRNPs. Non- snRNP proteins are also required for splicing and are likely to interact with other splicing components only transiently. Consistent with the fact that yeast contain fewer introns than mammals, their spliceosomes appear to be comprised of fewer proteins. One particular class of proteins found in metazoans that are not present in yeast is the SR family of proteins. These factors contain a number of serine/argenine (SR) or argenine/serine (RS) dipeptide repeats believed to participate in protein-protein interactions (Zahler et al., 1992). These SR proteins are essential for splicing and play a role in 5’ SS selection (Horowitz and Krainer,
1994). Although there is not an obvious family of SR protein homologues in yeast, several snRNP-associated yeast splicing factors do contain SR-like domains, indicating that their functions may be conserved even if their manner of association with the spliceosome is not.
1.3.2 snRNAs and snRNPs
Among the five snRNAs and the snRNP particles they form, U1, U2, U4 and U5 are similar. The U1-U5 snRNAs are modified by a unique trimethylguanosine cap structure that is not found in the U6 snRNA. Similarly,
5 only U1, U2, U4, and U5 snRNAs contain a conserved Sm site, a structural domain that allows binding to the seven common Sm core proteins, B, D1, D2,
D3, E, F and G (Branlant et al., 1982; Mattaj et al., 1986). The human Sm proteins were initially identified by their ability to cross-react with antisera from patients with the autoimmune disorder systemic lupus erythematosus and were subsequently named based on their mobilities during gel electrophoresis (Lerner and Steitz, 1979; van Venrooij, 1987). These proteins are known to initiate snRNP assembly in the cytosol by associating with the conserved Sm site in the snRNAs upon their export from the nucleus (Branlant et al., 1982; Mattaj and De
Robertis, 1985). The association of the Sm proteins then allows hyper- methylation of the 5’ cap of the snRNAs which together with the Sm proteins provides a signal for import into the nucleus (Fischer and Luhrmann, 1990;
Hamm et al., 1990). After returning to the nucleus, the remainder of the snRNP specific proteins associate with the particle to complete the maturation process
(Zieve and Sauterer, 1990). The U6 snRNP is distinct in that it contains a core of
LSm (for like-Sm) proteins, resulting in a similar overall structure (Mayes et al.,
1999). There are also differences between the yeast and human snRNAs.
6 Although the sizes of the U4, U5 and U6 snRNAs are similar in both species, the
U1 and U2 snRNAs in yeast are much larger than their human counterparts
(Ares, 1986).
The large majority of introns are called U2-dependent introns and are spliced by the major spliceosomal snRNPs U1, U2, U4, U5 and U6. Interestingly, a minor class of unique introns have been identified in some more advanced organisms including Arabadopsis, Drosophila, Xenopus, mice and humans. This rare class of introns are called U12-dependent and are spliced by a group of less abundant snRNPs including U11, U12, U4atac and U6atac in coordination with the standard U5 snRNP. This class of introns was first identified due to their unusual intron termini that contained an AT-AC sequence instead of the highly conserved GT-AG sequence (Burge, 1999). It was later found that the 5’ and branchpoint sequences in this class of introns were complimentary to the U11 and U12 snRNAs, two low-abundance snRNAs with previously unknown function. The involvement of these two snRNAs in splicing was supported by evidence of interactions between U11, U12, and U5 (Hall and Padgett, 1996;
Tarn and Steitz, 1996b). Two additional novel snRNA components, U4atac and
U6atac, which appear functionally analogous to U4 and U6 in the standard U2-
7 type spliceosome, were also identified (Tarn and Steitz, 1996a). The origins and possible advantages of these two systems are currently unclear.
1.3.2.1 The U1 snRNP
During the nearly three decades of splicing research, substantial effort has focused on understanding the U1 snRNP and its role in 5’ SS selection, primarily due to the fact that the binding of this particle to the pre-mRNA not only commits it to the splicing pathway but also plays a crucial role in identifying the 5’ site of cleavage. Despite the conserved and critical function of this particle, the U1 snRNPs found in mammalian and yeast systems vary greatly. The first difference lies in their snRNA molecules. Surprisingly, the yeast U1 snRNA is much larger than its mammalian counterpart (figure 1.4). While there is strong sequence homology in the 5’ ends and the Sm binding sites, the yeast molecule lacks a 3’ terminal stem/loop element that is found in the human U1 snRNA.
Included in the conserved regions is the 5’ stem/loop I, a structure that likely serves as a binding site for several snRNP-specific proteins (Guthrie and
Patterson, 1988). Much of the additional sequence in the yeast U1 snRNA can
8 be deleted without impairing cell viability, suggesting that the molecule contains a non-essential domain (Siliciano et al., 1987; Siliciano et al., 1991).
The second major difference between the yeast and mammalian U1 snRNPs lies in their protein composition. The mammalian U1 particle contains the seven conserved Sm core proteins as well as three U1-specific proteins, U1-
A, U1-C and U1-70K (Hinterberger et al., 1983). The yeast U1 snRNP, however, contains at least ten U1-specific proteins. In addition to the Sm core proteins and homologues of the human U1-A (yeast Mud1p), U1-C and U1-70K (Snp1p), seven additional proteins have been identified: Snu71p, Snu56p, Prp42p,
Nam8p, Prp39p, Prp40p, and Luc7p (Fortes et al., 1999; Gottschalk et al., 1998;
McLean and Rymond, 1998; Neubauer et al., 1997; Rigaut et al., 1999).
Although there is no extensive sequence homology between these yeast-specific factors and any known mammalian factors, it has been suggested that functional mammalian counterparts are likely to exist (Fabrizio et al., 1994; Gottschalk et al., 1998). These mammalian proteins may be very weakly associated with the
U1 particle and therefore not recovered during biochemical fractionation, or that they may interact only transiently during specific stages in splicing. This organizational heterogeneity may be due to the innate differences between
9 transcript structure in yeast and mammalian systems. Yeast have very few introns that contain highly conserved splice site sequences while humans have many introns with less conserved signals. Mammalian systems may therefore require more factors to help distinguish the proper splicing signals of a given transcript, for example the SR family of proteins know to assist in 5’ SS selection.
As yeast introns are less variable, it is possible that the yeast proteins performing the analagous function are fewer in number and may be integral U1 components that need not be recruited differentially (Fabrizio et al., 1994; Gottschalk et al.,
1998).
1.3.2.2 The U1-C Protein
Despite the identification of the numerous proteins of the U1 snRNP, the functions of most of these proteins remain largely unknown. The main exception to this is the U1-C protein, the most extensively studied U1-specific factor. The human U1-C protein is needed for association of the U1 snRNP with the 5’ SS in mammalian transcripts (Heinrichs et al., 1990). The U1-C protein does not
contain any of the well-characterized RNA binding motifs but does have a C2H2- type zinc finger in its N-terminus. Deletions and mutations in this motif prevent
10 the human U1-C protein from binding to the U1 snRNP. In addition, the incorporation of U1-C into the U1 particle requires the U1 snRNA and one or more U1 proteins (Nelissen et al., 1991). More specifically, the N-terminal domain of U1-70K and the common snRNP Sm proteins are required for U1-C to bind to the U1 snRNP. Cross-links between U1-C and the SmB protein indicate that this is likely the core Sm factor required for U1-C association (Nelissen et al.,
1994). In a continued effort to understand its role in stabilizing the U1/5’ SS interaction, human U1 snRNPs were biochemically reconstituted with various mutant versions of the U1-C protein to show that the N-terminal domain of U1-C is sufficient to stimulate U1/5’ SS complex formation (Will et al., 1996). Although the precise role of the U1-C zinc finger domain is still unclear, this motif has been shown to bind zinc ions (Dumortier et al., 1998).
The yeast counterpart of the mammalian U1-C protein is Yhc1p. YHC1, the gene encoding Yhc1p, is essential for cell growth and the yeast U1-C protein also associates with the U1 snRNP in yeast. Depletion of yeast U1-C does not appear to affect U1 snRNP assembly but blocks pre-mRNA splicing in vitro, apparently by preventing formation of the commitment complex. Interestingly, the 5’ arm of the U1 snRNA that normally binds to the 5’ SS was found to be
11 hypersensitive to RNase digestion when U1-C was depleted, indicating that this protein may contribute to proper U1 snRNA structure (Tang et al., 1997).
Three lines of evidence suggest that the U1-C protein plays a critical role in the recognition or definition of the 5’ SS. First, three separate studies found that despite the well-established role for the base-pairing interaction between the
U1 snRNA and the 5’ SS in commitment complex formation, the correct splice site sequence is recognized even in the absence of the 5’ end of the U1 snRNA.
These findings indicate that other factors in addition to the U1 snRNA are important for the recognition of the 5’ SS (Du and Rosbash, 2001; Lund and
Kjems, 2002; Rossi et al., 1996). Second, subsequent studies revealed that the
U1-C protein is able to select and bind to the nucleotide sequence GUAU in the absence of the 5’ end of the U1 snRNA, indicating that this protein directly contacts the pre-mRNA to promote the binding of the U1 snRNP to the 5’ SS.
The authors of the study propose that this interaction may precede the U1 snRNA/5’ SS base pairing. This conclusion is based on the theory that the protein/RNA interaction is likely to be more stable and that RNA base pairing required higher temperatures, suggesting the need for a conformational change to allow this interaction (Du and Rosbash, 2002). Third, the human U1-C protein
12 has also recently been shown to be important for the selection of weak 5’ SS sequences through interactions with regulatory splicing factors including TIA-1
(Forch et al., 2002).
1.4 The Dynamic Nature of the Spliceosome
1.4.1 Assembly of the Spliceosome
Studies in both the yeast and mammalian systems revealed a cycle of spliceosome assembly and disassembly on a pre-mRNA (figure 1.5) (Cheng and
Abelson, 1987; Konarska and Sharp, 1986; Konarska and Sharp, 1987). The first step that commits a pre-mRNA to the splicing pathway is recognition of the
5’SS by the U1 snRNP, initiating the commitment complex (CC) in yeast and the early (E) complex in mammals. This initial step does not require energy (ATP) for formation. U1 is the only component that can bind in the absence of ATP, and its interaction with the pre-mRNA is in general required for the other snRNPs to bind. After the association of the U1 snRNP with the 5’SS, the U2 snRNP recognizes and binds to the branchpoint to form complex B in yeast, called A complex in mammals. Addition of the U2 snRNP is the first energy-dependent step in the splicing pathway, and binding of the U2 snRNP is mediated in part by
13 the U1 snRNP as well as additional non-snRNP factors that bridge the two components. After U2 snRNP binding, the U4/U6.U5 tri-snRNP particle joins complex B to form complex A2-1 in yeast or B1 in mammals. Within the tri- snRNP particle there is extensive base pairing between the U4 and U6 snRNAs that forms two stem structures, and the U5 snRNP appears to be associated through protein-protein interactions.
Once all five snRNPs are present, the complex undergoes a series of rearrangements. The interaction between the U1 snRNA and the 5’ SS is disrupted, and the U1 snRNP particle is released from the complex. Similarly, the base pairing between the U4 and U6 snRNAs is also disrupted, allowing the release of the U4 snRNP and new base-pairing to form between the U2 and U6 snRNAs. The release of U1 and U4 from the splicing complex appears to be simultaneous, resulting in complex A1 in yeast or B2 in mammals that contains only U2, U5 and U6 snRNPs. At this stage, the complex undergoes structural rearrangements to form complex A2-2 in yeast or C1 in mammals. The spliceosome is then activated by another conformational change and is designated as complex A2-3 in yeast or C2 in mammals. Following catalysis, the mature mRNA is released, leaving complex I that contains the U2, U5 and U6
14 snRNPs as well as the lariat intron. The complex then dissociates and the snRNPs are then recycled for future rounds of splicing.
1.4.2 DExD/H box Proteins
Although energy in the form of ATP is required for splicing, there is no energy requirement for the transesterification reactions that result in intron removal. The cycle of spliceosome assembly, however, involves numerous structural rearrangements that are likely to require energy. Many of the conformational changes during spliceosome assembly correspond to changes in
RNA/RNA interactions, reactions that often involve proteins belonging to the
DExD/H box family of ATPases. Consequently, the energy requirement for splicing has been attributed to the involvement of numerous DExD/H box proteins that are required for spliceosome assembly and activation.
The DExD/H box family is a conserved group of proteins found in almost all organisms ranging from bacteria to humans and are also present in many viruses. Proteins in this family are involved in essentially all RNA-related biochemical processes such as transcription, RNA editing, pre-mRNA splicing,
RNA export, ribosome biogenesis, translation, and mRNA decay (Tanner and
15 Linder, 2001). The DExD/H box proteins contain six to eight conserved motifs and are named for the amino acid sequence found in one of these highly conserved domains (figure 1.6) (Linder et al., 1989; Tanner and Linder, 2001).
Most of these proteins have been shown to possess NTP hydrolysis activity that either requires or is stimulated by a nucleic acid cofactor, and some have also demonstrated the ability to unwind RNA or DNA duplexes in vitro (Schwer, 2001).
Eukaryotic initiation factor 4A (eIF4A) was the first DEAD box protein shown to modify RNA structures in an ATP-dependent manner and was subsequently shown to resolve RNA duplexes in vitro (Ray et al., 1985; Rozen et al., 1990).
This finding in addition to a study showing that another DEAD box protein, p68, also demonstrated RNA unwinding activity led to the tentative assumption that all
DEAD box proteins were also RNA helicases (Hirling et al., 1989; Linder et al.,
1989).
This classification of DExD/H box proteins was further supported by comprehensive sequence analyses that revealed that conserved sequences in these apparent RNA helicases were also present in the well-characterized DNA helicases, leading to a systematic classification of all types of helicases into superfamilies based on sequence similarity. Most DExD/H box proteins are
16 grouped into superfamily 2, while many viral RNA helicases are found in superfamily 1 along with a subset of DNA helicases (Gorbalenya and Koonin,
1993).
The function of the conserved motifs found in DExD/H box proteins have been revealed by mutational analysis. A summary of the likely functions of each of the DExD/H motifs is provided in figure 1.7. Early sequence analysis led to the prediction that motif I (AXXXXGKT) was a typical sequence for the A-motif of
NTP binding proteins while motif II appeared to be a version of the B-motif of
NTP binding proteins (Fry et al., 1986; Walker et al., 1982). Altering the conserved domains within eukaryotic initiation factor 4A (eIF-4A) confirmed that motif I (AXXXXGKT) is indeed required for ATP binding and showed that motif VI
(HRIGRXXR) is involved in ATP hydrolysis. Motif II (DEAD) mutations also affected ATP hydrolysis and were additionally found uncouple the ATPase and
RNA helicase activities of eIF4A (Pause and Sonenberg, 1992). Motif III (SAT) is essential for RNA unwinding and like the DEAD motif, mutations in the SAT motif have been shown to dissociate NTP hydrolysis from RNA unwinding activity
(Pause and Sonenberg, 1992; Schwer and Meszaros, 2000). Studies of the vaccinia virus helicase NPH-II revealed a similar pattern and also indicated that
17 the histidine residue in the DExH motif of NPH-II is required for coupling this protein’s NTPase and helicase activities (Gross and Shuman, 1995).
The systematic classification of RNA and DNA helicases also allowed predictions to be made about the overall structures of these proteins. Crystal structures of a number of helicases have since become available and verify many of these predictions. The enzymatic cores of these helicases consist of two domains connected by a linker region. The conserved motifs face into a cleft that lies in between the domains, providing a site for substrate and NTP binding
(Tanner and Linder, 2001). Additionally, the crystal structure of the helicase domain of the hepatitis C virus NS3, a DExH RNA helicase, was recently solved and may help provide additional insights into helicase mechanism (Kim et al.,
1998; Yao et al., 1997). Crystal structure analysis has also provided information as to how some of the conserved motifs may function. For example, crystal structures indicate that motif I forms a pocket that can bind the NTP phosphates while motif II binds the ß and γ phosphates through Mg2+ and coordinates NTP hydrolysis with a water molecule (Tanner and Linder, 2001).
Although the central core regions of many DNA and RNA helicases appear very similar, these molecules do differ in several important ways. RNA
18 helicases often have large amino- or carboxy-terminal extensions that are not typically found in DNA helicases. These extensions are projected to have additional functions such as an additional enzymatic activity, nuclear signaling, or providing specificity through interactions with protein cofactors (Tanner and
Linder, 2001). RNA helicases also differ from many of the DNA helicases in terms of enzymatic processivity, likely reflecting their structural and mechanistic differences. The DNA helicases in superfamilies 3, 4, and 5 are more distantly related to the RNA helicases and appear to work as hexamers, forming doughnut-shaped structures that can be visualized by electron microscopy.
These helicases are proposed to processively unwind long stretches of duplexed
DNA by rolling around one of the strands of DNA that is threaded through its hollow center (Patel and Picha, 2000). The RNA and DNA helicases of superfamilies 1 and 2, on the other hand, appear to work as monomers or dimers and tend to be less processive in nature. Monomers have been proposed to function by the inchworm model. In this model, the distance between the domains of the helicase varies with NTP binding and hydrolysis while other regions of the proteins interact with the nucleic acid backbone. This allows the molecule to act as a rachet, “creeping” along the substrate like an inchworm.
19 Dimers may work by the active-rolling model. In this case, the two monomer subunits may have alternating conformations based on NTP binding and hydrolysis that provide different affinities for the double-stranded and single- stranded regions of the substrate. This would allow the complex to “roll” along the substrate to disrupt the double-stranded regions (Tanner and Linder, 2001).
The active rolling model may in part explain the lack of processivity seen with some molecules as the enzymes display stages of lower affinity for the substrate, providing the opportunity for the helicase to “fall off”.
To understand how DExD/H box proteins function mechanistically, it is critical to examine how they recognize and interact with their specific substrates.
In some cases, these proteins interact with their substrates in a structure- dependent manner. The E. coli DEAD box protein DbpA forms a specific interaction with a hairpin of its 23S rRNA substrate through its carboxyl-terminal domain (Tsu et al., 2001). However, many other DExD/H box proteins exhibit rather promiscuous in vitro RNA unwidinding activities, suggesting that they may not recognize their substrates based on nucleic acid sequence or structure alone
(Tanner and Linder, 2001). The substrate specificity of many DExD/H box proteins is likely determined by interactions between the non-conserved regions
20 of these factors and the proteins surrounding their nucleic acid substrate. For example, the amino-terminal regions of two splicing DExD/H box proteins,
Prp22p and Prp16p, have been shown to be necessary for these factors to localize to the spliceosome (Schneider and Schwer, 2001; Wang and Guthrie,
1998).
Several recent studies have indicated that proteins may actually constitute part of the helicase’s target. First, one study asked whether a DExD/H box helicase was capable of disrupting a protein-RNA interaction. The viral DExD/H box protein NPH-II was shown to unwind an RNA duplex bound by the U1-A protein, displacing the protein from the RNA (Jankowsky et al., 2001). Although it is unclear whether NPH-II directly promotes protein dissociation or whether U1-
A protein removal was an indirect result of the alteration of the RNA binding site due to RNA unwinding, this study shows that a helicase is able to act as an
RNPase to alter the interaction between an protein and an RNA.
Genetic evidence suggests that two splicing DExD/H proteins may also disrupt protein-RNA interactions. First, specific mutations altering the U1-C protein can eliminate the requirement for the DEAD box protein Prp28p, suggesting that Prp28p may counteract the effects of the U1-C protein, a factor
21 thought to stabilize the U1/5’SS interaction (Chen et al., 2001). Second, a similar situation appears to occur at the branchpoint sequence. The branchpoint is initially bound by the branchpoint binding protein (BBP), an interaction that is likely stabilized by Mud2p, a protein that binds to a region downstream of the branchpoint. These two proteins must be displaced to allow the U2 snRNA to bind to the branchpoint (Berglund et al., 1998; Berglund et al., 1997). The deletion of Mud2p can bypass the requirement for the putative helicase Sub2p, indicating that Sub2p may normally displace Mud2p during spliceosome assembly (Kistler and Guthrie, 2001). Taken together, these recent findings indicate that DExD/H box proteins may remodel RNPs.
Importantly, these findings may also provide, at least in part, an explanation as to why most putative RNA helicases do not demonstrate RNA unwinding activity in vitro. It is likely that under such conditions an isolated
DExD/H box protein may not have available protein cofactors necessary for recognizing its substrate. This possibility was alluded to in earlier studies; eIF4A’s in vitro unwinding ability was dependent on the addition of eIF4B, a protein predicted to contain and RNA binding motif (Rozen et al., 1990). Another
DEAD box protein involved in mRNA export, Dbp5p, was only found to have in
22 vitro unwinding activity when purified from cell extracts, indicating the co- purification of essential cofactor(s) (Schmitt et al., 1999, Tseng et al. 1999).
1.4.2.1 DExD/H Box Proteins in splicing
According to the prevailing model, pre-mRNA splicing in yeast requires at least eight DExD/H box proteins: Prp5p, Brr2p, Prp28p, Sub2p (yUAP56), Prp2p,
Prp16p, Prp22p and Prp43p (Staley and Guthrie, 1998). A diagram of the splicing cycle indicating the steps where each putative helicase is required is shown in figure 1.8. Although their specific targets and mechanisms of function remain unclear, the stages of spliceosome maturation requiring each factor have been delineated and putative RNA/RNA targets have been identified.
The first RNA/RNA duplex formed during commitment of a pre-mRNA to splicing is the base pairing between the 5’ end of the U1 snRNA and the 5’ SS.
Before the first catalytic step, this interaction is disrupted and replaced by a mutually exclusive interaction between the U6 snRNA and the 5’ SS (Nilsen,
1998). This rearrangement requires not only the unwinding of the U1 snRNA/5’
SS duplex but also the disruption of the extensive base pairing between the U4 and U6 snRNAs. Prp28p is required for the switch of U1 for U6 at the 5’ SS and
23 is thought to disrupt the early interaction between the U1 snRNP and the 5’ SS
(Staley and Guthrie, 1999). Prp28p may also disrupt the interaction between the
U1-C protein and the pre-mRNA, supporting the model of DExD/H box proteins as RNPases (Chen et al., 2001). The function of Prp28p at the 5’ SS appears to be coupled to the function of Brr2p, another DExD/H protein thought to disrupt the U4/U6 RNA duplex. This conclusion was reached as both Brr2p and it’s human homologue U5-200K demonstrate the ability to dissociate the U4 and U6 snRNAs in vitro (Laggerbauer et al., 1998; Raghunathan and Guthrie, 1998a).
The unwinding of U4 from U6 releases regions of the U6 snRNA required for ensuing interactions with the 5’ SS as well as the U2 snRNA. This rearrangement also releases the U4 snRNP, releasing it from the spliceosome at approximately the same time that the U1 snRNP is released (Yean and Lin,
1991).
The second step during spliceosome assembly is binding of the U2 snRNP to the branch point, an association that is the first energy-requiring step during splicing. The branch point sequence is initially recognized by the Mud2 protein (mammalian SF1). Two DExD/H box proteins, Prp5p and Sub2p
(mammalian UAP56), are required for U2 snRNP binding to the branchpoint,
24 likely explaining the energy requirement for this step. After initial recognition of the branch point by the branch point binding protein (BBP), the branch point sequence pairs with a portion of the U2 snRNA, much like the recognition of the
5’ SS by the U1 snRNA (Berglund et al., 1997; Nilsen, 1998). In a manner similar to Prp28p counteracting U1-C, Sub2p has been proposed to displace
Mud2p, a protein that initially binds to the branch point and must be displaced to allow binding of the U2 snRNP (Kistler and Guthrie, 2001). A switch between two competing U2 snRNA conformations also controls the binding ability of the
U2 snRNP (Zavanelli et al., 1994). Prp5p is thought to modulate an ATP- dependent conformational change in the U2 snRNA, possibly exposing the branch point recognition sequence within the U2 snRNA to facilitate binding to the branch point (O'Day et al., 1996; Ruby et al., 1993). Interestingly, the branch point recognition sequence is not sequestered by the secondary structure of the
U2 snRNA in its inactive conformation. It was noted, however, that mutations altering Prp9p, a protein that binds to the U2 snRNA, made the U2 branch point recognition site more accessible. It is therefore possible that similar to Prp28p and Sub2p, Prp5p may act as an RNPase and target the Prp9p/U2 snRNA association (Staley and Guthrie, 1998). In addition to its role in branch point
25 recognition, the U2 snRNA is also involved in interactions with the U6 snRNA that appear to be important for both catalytic steps of splicing. Interestingly, the interactions between U2 and U6 are also mutually exclusive with the initial pairing between U4 and U6. The role of the U2 snRNA at the catalytic core will be discussed below.
Following binding of U2 and the ensuing rearrangements that release U1 and U4 and allow pairing between U6 and U2, two additional DExD/H box proteins catalyze rearrangements that lead to spliceosome activation. The first is
Prp2p, a factor that interacts transiently with the spliceosome through Spp2p and is required to function before the first catalytic step (Kim and Lin, 1996; Roy et al., 1995). The second is Prp16p, a protein that also interacts with the spliceosome only transiently (Schwer and Guthrie, 1991). Prp16p is required after the release of the U4 snRNP for the second transesterification reaction and affects a rearrangement at the 3’ SS (Schwer and Guthrie, 1992). After both transesterification reactions have taken place, several factors are required for the disassembly of the spliceosome and recycling of factors for subsequent rounds of splicing. The DExD/H box protein Prp22p is involved in disassembly of the spliceosome, aiding in the release of the mature messenger RNA molecule
26 (Schwer and Gross, 1998; Wagner et al., 1998). The final DExD/H box factor assigned a role in the splicing cycle is Prp43p, a protein required for release of the lariat intron from the spliceosome (Arenas and Abelson, 1997; Martin et al.,
2002). After its release, the lariat intron is recognized by the debranching enzyme that recognizes the 2’-5’ bond and cleaves it, allowing rapid degradation of the now linear intron (Ruskin and Green, 1985). Finally, at least one splicing factor has been shown to be important for snRNP recycling. Prp24p has been shown to assist in the re-annealing the U4/U6 snRNAs, presumably providing a means for re-assembling U4/U6 particles (Raghunathan and Guthrie, 1998b).
1.4.3 The Holospliceosome Hypothesis
Although the cycle of interactions and rearrangements in the spliceosome has been carefully delineated, recent findings are challenging the traditional view of spliceosome assembly. Several studies have identified interactions in early spliceosomes that would not be observed under standard spliceosome assembly conditions. For example, interactions between the U4/U6/U5 tri-snRNP and the
5’ SS that occur independently of U2 snRNP binding to the branch point have been identified in both human and nematode systems (Konforti and Konarska,
27 1994; Maroney et al., 2000). Crosslinking experiments in HeLa extracts revealed that the U2 snRNA is associated with U4/U6 snRNA complexes (Wassarman and
Steitz, 1992). This finding is consistent with other data suggesting that an interaction between the U2 and U6 snRNAs is required for human U4 and U6 snRNAs to form stable complexes (Brow and Vidaver, 1995). When low- stringency purification conditions are used, the U2 snRNP is found to be part of the mammalian E complex (commitment complex in yeast) that was previously thought to include only the U1 snRNP (Das et al., 2000). Finally, a functional penta-snRNP complex containing all five snRNP particles as well as a large number of associated non-snRNP splicing factors was recently purified from yeast extracts (Stevens et al., 2002). These findings indicate that the previously determined stepwise assembly may reflect the dynamic nature of the spliceosome. It is likely that the components of the splicing machinery form a loose structure that may initially interact with a pre-mRNA as a whole; RNA and proteins within this complex may then undergo an ordered series of stable interactions. The biochemical fractionations previously used to define the
28 splicing machinery likely resulted in the removal of the factors that were not involved in stable interactions at the time of purification, leading to the stepwise assembly model.
1.4.4 Prp28p
The research presented in this work focuses on the study of Prp28p, one of the DExD/H box proteins required during splicing. PRP28 was among the first putative RNA helicase genes identified in yeast based on the presence of the conserved motifs of the DEAD box family (Chang et al., 1990). Mutational analysis of these conserved domains in Prp28p revealed extensive genetic interactions among these motifs, indicating that several of these regions are likely to be in close proximity and that their activities may be coordinated during the protein’s function (Chang et al., 1997). PRP28 is an essential gene and homologues have been identified in a number of organisms including humans
(Teigelkamp et al., 1997). A screen for pre-mRNA splicing mutants in yeast found that a cold-sensitive allele of PRP28, prp28-1, blocked the first catalytic step of splicing. prp28-1 is also synthetically lethal with a mutation in PRP24, the gene encoding a U6 snRNP binding protein, and that a mutation in PRP8, the
29 gene encoding a U5 snRNP protein, could suppress the prp28-1 allele. Based on these genetic interactions, it was proposed that Prp28p may unwind the
U4/U6 duplex during splicing (Strauss and Guthrie, 1991). Prp28p possesses
RNA-dependent ATPase activity but is unable to unwind RNA duplexes in vitro.
Initially, it was suggested that Prp28p was unlikely to be a tightly bound, integral snRNP protein. This conclusion was based on two findings. First, no detectable splicing snRNAs were found to co-precipitate with Prp28p. Second, splicing extracts depleted of Prp28p were rescued by the addition of purified Prp28p, indicating that no associated snRNP factors had been co-depleted during the removal of Prp28p (Strauss and Guthrie, 1994). In contrast, U5-100K, the human homologue of Prp28p that shares 37% identity, is a component of the U5 snRNP (Teigelkamp et al., 1997). Although yeast Prp28p does not co-purify with the U4/U6/U5 tri-snRNP, even under low salt conditions that allowed co- purification of several U1 and U2 components, other purification techniques show that Prp28p is present in yeast U5 snRNPs (Gottschalk et al., 1999; Stevens et al., 2001).
Although initial studies predicted that Prp28p may disrupt the U4/U6 snRNA interaction, later findings support a different role for this protein in
30 splicing. Work by Staley and Guthrie (1999) revealed that Prp28p’s role in splicing was likely to occur at the U1/5’ SS duplex. In these studies, the RNA-
RNA interaction between the U1 snRNA and the 5’ SS was artificially extended, providing a hyperstabilized duplex that resulted in a cold-sensitive splicing defect.
This defect could be suppressed by mutations in the U1 snRNA predicted to weaken the duplex while mutations in the U1 snRNA predicted to further strengthen the duplex proved lethal. Importantly, extending the base pairing between the U6 snRNA and the 5’ SS was able to relieve the splicing defect resulting from the extension of the U1/5’ SS interaction, indicating that the U6 snRNA is able to compete with the U1 snRNA for binding at the 5’ SS. The splicing defect due to this hyperstabilization was also exacerbated in a prp28-1 mutant extract, indicating that Prp28p may normally help to destabilize the U1/5’
SS duplex. Utilizing biochemical techniques to isolate intact spliceosomes from in vitro splicing reactions, it was shown that all snRNPs were present in stalled spliceosomes from both Prp28p-depleted extracts as well as in reactions containing hyperstabilized U1/5’ splice sites. When recombinant Prp28p was added to these in the presence of ATP, the U1 and U4 snRNPs were released.
Although these results do not directly prove that Prp28p is functioning at the
31 U1/5’ SS, they do indicate that this is a likely target. Additional support for this from the human system has also emerged. Recent studies revealed that the human U5-100K protein, the human counterpart of Prp28p, was found to cross- link to the 5’ SS through its ATPase site (TAT), placing the active site of this
DEAD box protein in direct contact with its proposed target (Ismaili et al., 2001).
In addition to direct evidence for Prp28p’s role at the 5’ SS, secondary support for this function comes from the finding that the U4/U6 snRNA duplex,
Prp28p’s initially proposed target, is likely resolved by another splicing DExD/H box protein, Brr2p. Studies of the human U5 snRNP revealed that two putative helicases cross-linked to the U5 snRNA, U5-100K (yeast Prp28p) and U5-200K
(yeast Brr2p). Due to their proximity to the U4/U6 snRNA duplex, it seemed likely that one of these putative helicases was involved in their dissociation. In order to delineate which protein was involved in this unwinding, purified U5 snRNPs that lacked either U5-100K (Prp28p) or U5-200K (Brr2p) were incubated with either deproteinized yeast U4/U6 snRNA hybrids or artificial RNA duplexes.
The ability to unwind these RNAs was found to depend on U5-200K (Brr2p), not
32 U5-100K (Prp28p) (Laggerbauer et al., 1998). This conclusion was supported by the finding that Brr2p in yeast was able to dissociate the U4 and U6 snRNAs in vitro (Raghunathan and Guthrie, 1998a).
1.4.5 Recognition of the 5’ Splice Site
The first entity to recognize the 5’ SS during spliceosome assembly is the
U1 snRNP. The involvement of the U1 snRNP in pre-mRNA splicing was first proposed based on the observation that the sequence of the 5’ end of the U1 snRNA was complementary to the conserved sequence at the 5’ SS (Lerner et al., 1980; Rogers and Wall, 1980). Subsequent experiments showed that the U1 snRNP binds to 5’ SS and that this interaction is a requirement during in vitro splicing (Kramer et al., 1984; Mount et al., 1983). Some compensatory base changes in both the human and yeast U1 snRNAs suppress mutations in the 5’
SS of the pre-mRNA, confirming the importance of this interaction (Seraphin et al., 1988; Siliciano and Guthrie, 1988; Zhuang and Weiner, 1986). Once the recognition of the 5’ SS by the U1 snRNP had been established, the timing this interaction during the splicing reaction was investigated. Although the first stable pre-mRNA/snRNP interaction initially detected by native gel analysis was an
33 interaction with the U2 snRNP, it was determined that proper base pairing between the U1 snRNA and the pre-mRNA was required for U2 snRNP association and splicing (Cheng and Abelson, 1987; Konarska and Sharp, 1986;
Lamond et al., 1988; Pikielny et al., 1986; Seraphin et al., 1988). Additionally, a commitment complex between the U1 snRNP and the 5’ SS can form in the absence of ATP and the U2 snRNP (Legrain et al., 1988; Ruby and Abelson,
1988). Definitive evidence for commitment to splicing via U1 binding came through a study using yeast extracts metabolically depleted of individual snRNPs.
These experiments showed that the U1/pre-mRNA complexes were stable, could occur in the absence of U2 and could be chased into functional spliceosomes
(Seraphin and Rosbash, 1989).
Interestingly, although some 5’ SS mutations could be suppressed by compensatory mutations in the U1 snRNA, others could not, indicating that the recognition of the 5’ SS is a very complex process and was unlikely to occur only by this RNA/RNA interaction. In addition, these non-suppressing mutations also appeared to affect 5’ SS specificity (Guthrie, 1991; Seraphin et al., 1988;
Seraphin and Rosbash, 1990; Siliciano and Guthrie, 1988; Zhuang and Weiner,
1986). These observations suggested that one or more factors in addition to the
34 U1 snRNA also recognize these nucleotides at the 5’ SS. As predicted, additional snRNAs and proteins have also been shown to recognize the 5’ SS.
First, the U6 snRNA has also been found to play a critical role in 5’ SS recognition. The conserved ACAGA sequence within the U6 snRNA was found to cross-link to the 5’ SS in both yeast and mammalian systems (Sawa and
Abelson, 1992; Sontheimer and Steitz, 1993; Wassarman and Steitz, 1992).
Subsequent genetic suppression experiments confirmed that this U6 sequence forms Watson-Crick base pairing with the 5’ SS and influences splice site selection (Kandels-Lewis and Seraphin, 1993; Lesser and Guthrie, 1993).
Interestingly, the U6/5’ SS interaction is mutually exclusive with the U1/5’ SS interaction as overlapping regions of the 5’ SS are recognized by both snRNAs.
Although the interactions between the 5’ SS and U1 and U6 both occur before the first catalytic step, it was determined that the after the U1 snRNP binds to this region the interaction is disrupted and replaced by the U6/5’ SS duplex (Konforti et al., 1993). This conclusion is consistent with the speculations that the U6 snRNA is important for catalysis, a circumstance requiring that the U6 snRNA be positioned near the splice sites during the transesterification reactions (Yean et al., 2000).
35 The U5 snRNA has also been shown to play a role in 5’ SS selection.
Mutations in the U5 snRNA were found to activate the use of improper 5’ SS sequences when the normal 5’ SS had been altered (Cortes et al., 1993;
Newman and Norman, 1991; Newman and Norman, 1992). Cross-linking experiments also revealed interactions between the U5 snRNA and the 5’ and 3’ splice sites (Sontheimer and Steitz, 1993; Wyatt et al., 1992). Specifically, the
U5 snRNA can be cross-linked to the last nucleotide of the 5’ exon before and during catalysis. The U5 snRNA can also be cross-linked to the first nucleotide of the 3’ exon, but only after the first transesterification reaction (Newman et al.,
1995). These data led to the idea that the U5 snRNA functioned as a tether to retain the first exon after the first catalytic step and then align it with the second exon to allow the second catalytic step.
In addition to the multi-step recognition of the 5’ SS by the U1, U6 and U5 snRNAs, a number of proteins have also been implicated in assisting in 5’ SS recognition. The first proteins found to have such a role are the essential family of SR proteins found in metazoan systems. These proteins contain domains rich in arginine/serine dipeptides and were found to influence 5’ SS choice during alternative splicing. Some SR proteins stabilize the U1 snRNA/5’ SS interaction
36 and mediate interactions between the U1 and U2 snRNPs to aid in recognition of the splice sites (Fu and Maniatis, 1992; Kohtz et al., 1994; Zuo and Manley,
1994). In addition, high concentrations of purified SR proteins were able to compensate for the loss of the U1 snRNP during in vitro splicing reactions
(Crispino et al., 1994; Tarn and Steitz, 1994). Since this finding, a number of other proteins have also been shown to aid in recognition of the 5’ SS. One study found that eight different proteins could be cross-linked to the 5’ SS region of the pre-mRNA in the yeast commitment complex. These factors include yCBP80, a component of the cap binding complex, and seven U1 snRNP proteins including U1-C, U1-70K, Nam8p, Snu56p, SmD1, SmD3 and SmB
(Zhang and Rosbash, 1999). The finding that U1-C was among these proteins is not surprising, as it has since been well established that this protein plays an important role in commitment complex formation. It was not unexpected that
Nam8p was also identified, as Nam8p may aid in 5’ SS selection during splicing of MER2, a meiosis specific transcript containing a weak 5’ SS sequence
(Nakagawa and Ogawa, 1997). The involvement of the three Sm proteins in commitment complex formation has also been elucidated. The Sm proteins D1,
D3 and B have highly charged C-terminal tails, the deletion of which prevents the
37 stable association of U1 with the 5’ SS (Zhang et al., 2001). The identification of yCBP80 is also not surprising, as interactions between the cap binding complex and the U1 snRNP are know to be important during commitment complex formation, specifically for assisting the U1 snRNP in binding to the cap-proximal
5’ SS (Lewis et al., 1996). In addition to the proteins found to cross-link to the 5’
SS during commitment complex formation, a recent study found that Prp8p, a conserved component of the U5 snRNP, may play a direct role in recognizing the
5’ SS early in spliceosome formation (Maroney et al., 2000). The following section contains a detailed discussion of Prp8p and its proposed role at the catalytic core of the spliceosome.
1.4.6 Prp8p and the Catalytic Core of the Spliceosome
The catalytic core of the spliceosome is thought to form through the interaction of the U6 snRNA with the 5’ SS and the U2 snRNA, allowing the 2’ hydroxyl of the branchpoint adenosine to be juxtaposed to the conserved G at the 5’ SS to allow the first transesterification reaction. This model has developed through numerous genetic and cross-linking studies revealing extensive interactions between the U2 and U6 snRNAs (Villa et al., 2002). The U6 snRNA
38 is proposed to be the actual catalytic entity, indicating that the spliceosome may be a true ribozyme. Most RNA enzymes require metal ions as cofactors for catalysis, and it was recently determined that specific bases in the U6 snRNA bind Mg2+ ions, a strong indication that this molecule may indeed be the catalytic agent (Collins and Guthrie, 2000; Yean et al., 2000). A protein-free complex of the U2 and U6 snRNAs is able to bind a small RNA and activate the attack of a branch point adenosine on a catalytically important region of U6. This reaction is similar to the first step of splicing, further supporting the potential catalytic activity of the U2 and U6 snRNAs (Valadkhan and Manley, 2001).
In addition to the U6 and U2 snRNA molecules, Prp8p has also been proposed to play a critical role at the catalytic core of the spliceosome. Prp8p is a component of the U5 snRNP in both yeast and mammalian systems and is the most highly conserved splicing factor among species. PRP8 was first isolated as
RNA8 in a genetic screen seeking factors involved in pre-mRNA splicing in yeast and was found to encode an extremely large 260 kilodalton (Kd) protein that is associated with the U5 snRNA (Lossky et al., 1987; Lustig et al., 1986). PRP8 is an essential gene, and a yeast strain containing a temperature-sensitive prp8-1 allele is defective in splicing at high temperatures. Splicing extracts
39 immunodepleted of Prp8p are also defective in splicing, confirming a role for
Prp8p in pre-mRNA splicing (Jackson et al., 1988). The same antibody used for immunodepletion was subsequently employed to demonstrate that Prp8p is a component of the U5 snRNP, is present in the spliceosome throughout the splicing reactions and is also found in a post-splicing complex containing the excised lariat intron (Whittaker et al., 1990). Three separate investigations also took advantage of the antibody to the yeast Prp8p to identify the human homologue of this factor, hPrp8p. Two studies used the antibody to directly search for related human proteins and identified a protein of approximately 220
Kd that was a component of the U5 snRNP, providing the first evidence of a splicing factor conserved in yeast and humans (Anderson et al., 1989; Pinto and
Steitz, 1989). The third study sought factors that directly interact with the pre- mRNA, and in doing so identified a 220 Kd protein that cross-linked to the pre- mRNA. This protein also cross-reacted with the anti-Prp8p antibody, revealing their relatedness (Garcia-Blanco et al., 1990). This result in the human system was also confirmed by the finding that yeast Prp8p can also be cross-linked to the pre-mRNA during in vitro splicing reactions (Whittaker and Beggs, 1991).
40 A number of genetic and biochemical approaches have been employed in attempts to unveil the function of Prp8p during splicing. A mutation in DED1, a gene encoding a DExD/H box protein, was found to suppress the temperature- sensitivity of the prp8-1 mutation (Jamieson et al., 1991). Interestingly, subsequent experiments have suggested a role for DED1 in translation (Chuang et al., 1997). A mutation in PRP8 was found to suppress the cold-sensitive splicing defect caused by the prp28-1 mutation, and a novel allele of PRP8 was also identified in a screen for cold-sensitive splicing factors (Noble and Guthrie,
1996; Strauss and Guthrie, 1991). A combination of genetic and biochemical techniques revealed that inhibition of Prp8p reduced the ability of U5 to associate with U4/U6 and also inhibited the association of the U4/U6 and U5 snRNPs with the spliceosome (Brown and Beggs, 1992).
Prp8p appears to play important roles at both the 5’ and 3’ splice sites, providing the first evidence that Prp8p may be a component of the catalytic core of the spliceosome. Consistent with the finding that Prp8p could be cross-linked to the pre-mRNA, Prp8p was found to specifically interact with a large portions of the exons near the 5’ and 3’ splice sites as well as with a part of the polypyrimidine tract upstream of the 3’ SS in the intron (Teigelkamp et al., 1995a;
41 Teigelkamp et al., 1995b; Whittaker and Beggs, 1991). The pattern of contacts between Prp8p and the exons of the pre-mRNA closely mirrors the interaction between the U5 snRNA and the same regions of the transcript (Newman and
Norman, 1991; Newman and Norman, 1992; Sontheimer and Steitz, 1993; Wyatt et al., 1992). This observation led to the proposal that Prp8p may stabilize the interaction between the U5 snRNA and the pre-mRNA, assisting the U5 snRNA in exon alignment during splicing (Teigelkamp et al., 1995a). This conclusion was verified, at least in part, by the finding of extensive interactions between the
U5 snRNA and Prp8p (Dix et al., 1998; Urlaub et al., 2000). Interestingly it was later found that these regions of the U5 snRNA are not essential for viability, introducing the possibility that the role of the U5 snRNA at the exons is to present
Prp8p to the catalytic core (Collins and Guthrie, 1999).
In an extension of earlier cross-linking studies, it was determined that the
C-terminal portion of Prp8p (amino acids 1894-1898) specifically cross-links to the conserved GU dinucleotide at the 5’ SS (Reyes et al., 1999; Reyes et al.,
1996). This finding spurred genetic analysis revealing a large number of C- terminal alterations in Prp8p that suppress 5’ and 3’ splice site mutations by affecting the efficiency of the second catalytic step (Siatecka et al., 1999). The
42 initial finding of the cross-link between Prp8p and the 5’ SS may also have additional meaning aside from the possible catalytic function. A recent study found that Prp8p was able to cross-link to the 5’ SS during an early, U1 snRNP- independent, interaction between the tri-snRNP and the pre-mRNA. This indicates a possible role for this protein in early recognition of the 5’ SS and/or recruitment of the tri-snRNP to the spliceosome (Maroney et al., 2000).
An allele of PRP8, prp8-101, was isolated in a screen for factors required for recognition of the polypyrimidine tract and 3’ SS. Consistent with the results from the aforementioned studies, this report also demonstrated that Prp8p could be cross-linked to the 3’ SS (Umen and Guthrie, 1995a). In addition, it was found that the prp8-101 mutation prevented Prp8p from cross-linking to the 3’ SS
(Umen and Guthrie, 1995b). Extensive mutagenesis of PRP8 produced a large number of mutations that appeared to behave like prp8-101, and detailed analysis of these alleles allowed the definition of two classes of mutants. The first affected 3’ SS fidelity while the second affected 3’ SS selection.
Interestingly, these two classes of alterations mapped to unique domains within
43 Prp8p but in general mapped to the C-terminal portion of the gene, the same region of the protein implicated in functions at the 5’ SS (Umen and Guthrie,
1996).
Another class of PRP8 mutants appear to suppress defects in exon ligation caused by mutations at either the 5’ or 3’ splice sites. The alleles identified as 3’ SS selection mutants in the study by Umen and Guthrie (1996) were re-classified as general splice site suppressors in this study. Interestingly,
PRP8 mutants can also suppress a mutation in the ACAGA motif of the U6 snRNA, indicating a role in positioning the U6 snRNA and splice sites for catalysis (Collins and Guthrie, 1999). This hypothesis is supported by a number of additional findings. First, Prp8p was found to cross-link to position U54 of the
U6 snRNA. This residue is positioned between the region of the U6 snRNA that base pairs with the 5’ SS and the region that interacts with the U2 snRNA, likely forming the catalytic center of the spliceosome (Vidal et al., 1999). Second, an allele of PRP8, slt-21/prp8-21, was identified in a screen for mutations that are synthetically lethal with mutations in the U2 snRNA. Interestingly, this allele was also synthetically lethal with mutations in the U5 snRNA, providing a potential link
44 between Prp8p’s putative role in exon alignment and its interaction with the RNA components of the catalytic core (Xu et al., 1998).
In addition to the extensive volume of data that implicate Prp8p in assimilating numerous molecules at the catalytic core, Prp8p may also act as a regulator of the spliceosomal rearrangements that allow activation. A mutation in
PRP8, prp8-201, was identified as a suppressor of the U4-cs1 mutation. U4-cs1 extends the base pairing between the U4 and U6 snRNAs into the ACAGA box of the U6 snRNA, preventing its interaction with the 5’ SS. This led to the proposal that Prp8p may trigger the unwinding of the U4 and U6 snRNAs only after correct recognition of the 5’ SS by the ACAGA box of U6. Like the many other PRP8 mutations, this alteration to Prp8p maps to the C-terminal domain of the protein
(Kuhn et al., 1999). A large number of secondary suppressors of U4-cs1 were also identified and map to five domains throughout Prp8p. Genetic interactions between these domains indicate a possible intra-molecular fold within Prp8p
(Kuhn and Brow, 2000). A diagram showing the putative regulatory regions within Prp8p is shown in figure 1.9. Additional genetic interactions indicate that
Prp8p may regulate Brr2p, the putative U4/U6 helicase, and Prp28p, the putative
U1/5’ SS helicase, in order to control the rearrangements they promote to allow
45 spliceosome activation (Kuhn and Brow, 2000; Kuhn et al., 2002). Further support for these findings was provided upon the detection of a physical interaction between Prp8p and Brr2p (van Nues and Beggs, 2001). Although the evidence for a role for Prp8p in regulating Prp28p is not thoroughly convincing, the data presented in this dissertation provides additional support for this idea.
Interestingly, recent work has indicated that Prp8p as well as a number of other proteins may play a role in signaling downstream processing events that follow splicing. Prp8p is one of several proteins deposited in a complex at the exon-exon junctions of mRNAs following splicing, indicating a potential function in linking the splicing process to future events such as mRNA export (Le Hir et al., 2000a; Le Hir et al., 2000b). Finally, it is also of interest to note that studies of Prp8p appear to be relevant to the process causing the degenerative eye disease retinitis pigmentosa (RP). Mutations in human PRP8 have been identified in a number of families affected by an autosomal dominant form of the disease (McKie et al., 2001). Additional studies found that mutations in another splicing factor, PRP31, are also linked to autosomal dominant RP. Prp31p appears to tether U5 to the U4/U6 snRNPs, indicating that this process may be defective in individuals with RP (Makarova et al., 2002).
46 1.5 Goal of This Work
Although the role of DExD/H box proteins in pre-mRNA splicing has been well established, an understanding of the in vivo targets of these proteins and their mechanism of function is lacking. One such factor is Prp28p, an essential splicing factor thought to unwind the U1 snRNA/5’ SS duplex during spliceosome formation. The goal of this work is to continue studies of Prp28p in order to elucidate the components of its in vivo substrate and to better understand its role in splicing. The data presented here demonstrate that Prp28p may counteract the stabilizing effects of a number of U1 snRNP proteins including U1-C, Prp42p and Snu71p. These findings indicate that this DEAD box protein may function as an RNPase, working to rearrange protein and RNA interactions within the U1 snRNP to destabilize its interaction with the 5’ SS. Additional studies show that
Prp8p may play an important role in assisting Prp28p during the displacement of the U1 snRNP.
47 Figure 1.1. Selected types of alternative splicing. Exons are shown as open boxes, alternatively spliced exons are represented by hatched boxes. Splicing between the exons is indicated by v-shaped lines. A) constitutive splicing, B) exon inclusion/exon skipping, C) 5’ splice site choice, and D) splice site/no splice site. Adapted from Horowitz and Krainer, 1994.
48 A
B
C
D
49 A
Exon 1 AG GUAUGU UACUAAC YnAG G Exon 2
B
AG GURAGU YNYURAY YnAG G Exon 1 Exon 2
Figure 1.2. Consensus sequences of splicing signals. Shown are the conserved sequences found at the 5’ splice site, the branch site and the 3’ splice site in A) yeast introns and B) mammalian introns (R=purine, Y=pyrimidine, N=any nucleotide). Conserved adenosines at the branch site are shown in bold. Adapted from Horowitz and Krainer, 1994.
50 A A
B OH A
C + A
Figure 1.3. The chemical mechanism of pre-mRNA splicing. Intron removal occurs by two sequential transesterification reactions. A) The conserved adenosine at the branch point serves as the nucleophile, attacking the phosphodiester bond at the 5’ SS . B) This results in a lariat intron-exon 2 intermediate and a free exon 1. The hydroxyl group at the 5’ SS of exon one is the nucleophile for the second reaction, attacking the phosophodiester bond at the 3’ SS and resulting in ligated exons and a free lariat intron (C).
51 Figure 1.4. Secondary structures of the human and yeast U1 snRNAs. A) The yeast U1 snRNA is much larger than B) the human U1 snRNA. Boxes indicate the conserved domain. Adapted from Tang and Rosbash, 1996.
52 53 Figure 1.5. The spliceosome cycle. Spliceosomes assemble on pre-mRNA transcripts by sequential binding of the snRNPs and are then activated by a series of rearrangements. Following catalysis, the components are released and snRNPs are recycled for subsequent rounds of splicing. The names of each complex are noted according to the yeast terminology; the mammalian complex notation is provided in parentheses.
54 CC (E) U1 B (A)
U1 U2 U1 U2
Complex I A2-1 (B1)
U5 U2 U6 U1 U2 U6
55 U6 U4 U6 U4 U5 U4 U5
U1 A2-3 (C2)
U6 U4 U2 A1 (B2) U5 A2-2 (C1)
U6 U2
U6 U2 U5
U5 Figure 1.6. Conserved sequence motifs of DExD/H box proteins. A) Generalized positions of the conserved motifs are shown as colored boxes. B) The conserved amino acid sequences for the DEAD box family are shown. Upper case letters represent amino acids conserved at least 80% of the time while lower case letters represent amino acids conserved 50%-79% of the time. Adapted from Tanner and Linder, 2001.
56 A
I Ia Ib II III IV V VI 57
B
Motif I Motif Ia Motif Ib Motif II Motif III Motif IV Motif V Motif VI
A-TGTGKT PTRELa-Q TPGRl VlDEaD-m SAT LiF—T LvaTdvaaRGlD Y-HRiGRTgR-G S S S S
Motif Known or Suggested Function
I P-loop; Walker A NTP-binding motif; binds phosphates of NTP
Ia Binds substrate through sugar-phosphate backbone
Ib Substrate binding; not as highly conserved and may not always be present
II Walker B NTP-binding motif; binds ß and γ phosphates through Mg2+; coordinates hydrolysis of NTP with water molecule
III Binds γ phosphate; links NTP hydrolysis with unwinding activity
IV Substrate binding; known as IVa in SF1 DNA helicases
V Binds substrate through sugar-phosphate backbone; may interact with NTP
VI Binds γ phosphate; converts NTP binding/hydrolysis with domain 1 and 2 movement
Figure 1.7. Summary of known or suggested functions of DExD/H box protein motifs. Adapted from Tanner and Linder, 2001.
58 Figure 1.8. DExD/H box proteins in pre-mRNA splicing. The names of the proteins are positioned at the stages of spliceosome rearrangement where they participate.
59 U1
U1 U2 U1 U2 Prp5p Sub2p
U5 U2 U6 U1 U2 U6 U6 60 U4 U6 U4 U5 U4 U5 Prp28p Prp43p Prp22p U1 Brr2p U6 U4 U2 U5
U6 U2 Prp2p Prp16p U6 U2 U5
U5 Figure 1.9. Map of proposed functional domains within Prp8p. The numbers at the top of the diagram represent amino acid positions. The small red box represents a domain identified as important for 3’ splice-site fidelity. The large blue box represents the region containing mutations that can suppress splice site mutations. The black line at position 1861 represents the mutation found to suppress the U4-cs1 allele; white boxes represent the positions of the suppressors of this allele identified in a secondary screen. The lightning bolt indicates the position of the cross-link between the hPrp8 protein and the 5’ splice site. Black brackets indicate putative intra-molecular folds within the protein.
61 1964-1968
1 236 362 611 684 788 861 1094 1197 1371 1608 1817 1861 2065 2413 62
Prp8p CHAPTER 2
MATERIALS AND METHODS
2.1 Primer Extension
Total RNA was prepared from strains YTC105 (PRP28 YHC1), YTC102
(prp28::HIS3 prp28-102/TRP1) and YTC300 (prp28∆ YHC1-1). Strains were
grown at 30˚C to an approximate OD600 of 0.5 at which time half of the culture was shifted to 16˚C for 16 hours. Cells were collected by brief centrifugation and total RNA was extracted using a standard hot phenol/chloroform (pH 5.3) extraction. A total of 50 µg of total RNA was annealed with 50 fmol of 32P end- labeled actin primer (ACT4, 5’ tgccagatcttttccatatcgtccagttg 3’) in annealing buffer [150 mM KCl, 12 mM Tris-Cl (pH 8.0), 0.12 mM EDTA (pH 8.0)]. After annealing, 50 µl of RT solution [1 mM each dNTP in AMV RT buffer supplied by
Pharmacia (50 mM Tris-HCl (pH 8.3) 8 mM MgCl2, 50 mM NaCl and 1 mM DTT)] and 1 µl (15U) of AMV reverse transcriptase (Pharmacia) were added. Reverse
63 transcription was carried out at 45˚C for 1 hour after which RT products were purified on MicroSpin G-25 sephadex columns (Pharmacia) to remove excess 32P and salts. After products were dried using a speed-vac they were resuspended in 3 µl sequencing loading dye (1 mg/mL bromophenol blue and xylene cyanol in formamide). Products were separated by polyacrylamide gel electrophoresis on an 8 M urea 6% polyacrylamide (19:1) gel and visualized by autoradiography.
2.2 In vitro Splicing
Splicing extracts were prepared according to the liquid nitrogen grinding procedure (Umen and Guthrie, 1995). Liquid cultures were grown to an
approximate OD600 of 2.5-3 and then harvested by centrifugation at 3,304 g
(4,500 rpm in a Sorvall GS-3 rotor) for 10 minutes. Cells were washed once with
buffer AGK [10mM HEPES (pH 7.9), 1.5 mM MgCl2, 200 mM KCl, 10% glycerol, and 0.5 mM DTT] and centrifuged as above. The cell pellet was then resuspended in 0.4 volumes of buffer AGK containing one Complete mini protease inhibitor tablet (Roche) per 10 mLs of buffer and frozen in liquid nitrogen. The frozen cells were then ground to a fine powder using a mortar and pestle that was cooled with liquid nitrogen. After grinding the powder was quickly
64 thawed at room temperature and then kept on ice. The cell lysate was then centrifuged at 34,858 g (17,000 rpm in a Sorvall SS-34 rotor) at 4˚C for 30 minutes. The supernatant was subsequently subjected to ultra-centrifugation at
99,058 g (38,000 rpm in a 70.1 Ti rotor) at 4˚C for one hour. Approximately two- thirds of the supernatant was removed and dialyzed twice against two liters of buffer D [20 mM HEPES (pH 7.9), 50mM KCl, 2 mM EDTA, 20% glycerol, 0.5 mM DTT] for 1.5 hours each time. The extract was then centrifuged at 16.1 g in a microcentrifuge at 4˚C for one minute to pellet precipitates. The supernatant was moved to a fresh tube, frozen in liquid nitrogen and stored at –80˚C.
In vitro splicing was conducted in 10 µl reactions containing 4 µl splicing extract and 60,000 cpm (6 fmol) of 32P labeled actin pre-mRNA reporter transcript
in splicing buffer [2.5 mM MgCl2,1 mM spermidine, 2 mM ATP, 60 mM KPO4 (pH
7.0) and 3% PEG8000]. For reactions containing recombinant His6-Prp28p (see below), 0.4 µg (6 pmol) of recombinant protein was added to the reactions which were then pre-incubated without ATP at 25˚C for 15 minutes. After the mixture had been chilled on ice, ATP was added. All reactions were assembled on ice and then incubated at 25˚C for 20 minutes or 16˚C for one hour. Following this incubation, the reactions were briefly chilled on ice and then subjected to
65 phenol/chloroform (pH 5.3) extraction and ethanol precipitation. Precipitated splicing products were washed with 70% ethanol, dried and resuspended in 6 µl of water and 4 µl of loading dye (1 mg/mL each xylene cyanol and bromophenol blue in formamide). Splicing products (5 µl) were separated by polyacrylamide gel electrophoresis on an 8 M urea 8% acrylamide (29:1) gel and visualized by autoradiography.
2.3 Purification of His6-Prp28p and Production of anti-Prp28p Antibody
Plasmid pCA8065 [PRP28/pRSET-A (Invitrogen)] was transformed into
- λ bacterial strain BL21 (DE3) [F dcm ompT hsdS(rB- mB-) gal (DE3)]. Single colonies were inoculated into LB-ampicillin cultures and grown at 37˚C to reach
β an OD600 of 0.5 at which time isopropyl -thiogalactopyranoside (IPTG,
Invitrogen) was added to a final concentration of 0.2 mM to induce expression of
the recombinant His6-Prp28 protein. After three hours of induction, cells from 8 liters of cultures were collected by centrifugation at 3,304 g (4,500 rpm in a
Sorvall GS-3 rotor) for 10 minutes and washed in 50 mL of 50 mM Tris-Cl (pH
7.5), 1 mM EDTA for every liter of culture used. Cells were centrifuged as above and pellets were resuspended in 5 mL/g of cells in buffer A [40 mM HEPES-
66 NaOH (pH7.5), 1 mM EDTA, 0.5 mM DTT, 10% glycerol and one Complete protease inhibitor tablet per 50 mL of buffer (Roche)] containing 100 mM NaCl
(hereafter buffer A100). Lysozyme (Sigma) was added to 0.2 mg/mL and cells were incubated on ice for one hour. Cells were lysed using three cycles of sonication/freeze–thaw. Each cycle consisted of three 10-second bursts of sonication at a medium intensity setting followed by freezing in liquid nitrogen and rapid thawing in a 37˚C water bath. The crude lysate was centrifuged at
39,079 g (18,000 rpm in a Sorvall SS-34 rotor) at 4˚C for 30 minutes to remove cell debris. Polyethyleneamine (Sigma) was added to the supernatant to 0.25% with constant stirring for 1 hour at 4˚C to precipitate nucleic acids. Precipitates were removed by centrifugation at 39,079 g (18,000 rpm in an SS-34 rotor) at
4˚C for 30 minutes. Proteins in the supernatant were subjected to precipitation using ammonium sulfate (Sigma) at 40% saturation with constant stirring at 4˚C for one hour. Precipitated proteins were collected by centrifugation at 39,079 g
(18,000 rpm in a Sorvall SS-34 rotor) at 4˚C for 30 minutes, resuspended in 10 mL of buffer A100 and dialyzed against two changes of buffer A100 at 4˚C overnight. The protein suspension was then applied to a 6 mL heparin- sepharose CL-6B (Pharmacia) column that had been prepared by washing with
67 ten volumes of buffer A100 using a peristaltic pump with a flow rate of approximately 2 mL/min. After the protein solution was applied to the column by gravity flow, it was washed with approximately ten volumes of buffer A100 at 2 mL/min. Protein was then eluted using 15-mL step fractions of buffer A
containing 200, 300, 400, and 500 mM NaCl. His6-Prp28p is predominantly found in fractions A300, A400 and A500. These fractions were then dialyzed into buffer A50 containing only 1 mM EDTA and then applied to a 5-mL ProBond
Nickel resin (Invitrogen) column that was prepared by washing with approximately ten volumes of buffer A50/1 mM EDTA. After applying the combined dialyzed fractions to the column by gravity flow, the column was washed with 10 bed volumes each of A50, A100, A150, A300 and A50, all
containing 1 mM EDTA. The His6-Prp28 protein was then eluted with a 10 mM to
300 mM linear gradient of imidazole (Sigma) in buffer A50/1 mM EDTA.
Fractions containing His6-Prp28p were pooled and dialyzed into buffer D containing 50% glycerol before storage at 0.4 µg/µl (6 pmol/µl) at either –20˚C or
–80˚C.
Purified His6-Prp28p was submitted to Cocalico Biologicals, Inc.
(Reamstown, PA) for the production of an anti-His6-Prp28p antibody from rabbits.
68 After each test bleed the rabbit serum was tested using Western blots of yeast total protein extracts according to standard procedures (Sambrook, 1989). The final antibody was obtained after six boosts.
2.4 Testing U1 snRNA Mutations for Bypass of PRP28
Plasmids carrying mutant versions of the SNR19 gene that encode the U1 snRNA (U1-C4A, C4U, U5G, U5A, U5C and C8U) (Siliciano and Guthrie, 1988) were transformed into the yeast strain YTC65 [prp28::HIS3 pCA8032
(PRP28/URA3)]. Transformants were grown to saturation in SD liquid medium
and O.5 OD600 units of cells were collected. Cells were resuspended in 100µl of water and three ten-fold serial dilutions were performed. Spots of 4 µl were placed on plates containing 1 mg/mL 5-Fluoorotic acid (5-FOA, Toronto
Research Chemicals, Inc.) and incubated at 30˚C for five days.
2.5 Testing the Growth Phenotypes of U1-C L-13 Variants
Plasmids carrying mutant alleles of YHC1 (see table 2.2) were individually transformed into YTC65 [prp28::HIS3 pCA8032 (PRP28/URA3)] and YTC510
[yhc1::HIS3 pYHC1023 (YHC1-HA/URA3)]. Transformants were streaked to
69 plates containing 1 mg/mL 5-FOA to eliminate pCA8032 (PRP28/URA3) or pYHC1023 (YHC1-HA), respectively. The resulting strains were then grown in
SD-Leu liquid cultures to saturation after which 0.5 OD600 units of cells were collected. Cells were resuspended in 100 µl of water and three ten-fold serial dilutions were performed. From each dilution 4 µl of cells were spotted to plates that were incubated at various temperatures until growth in the least dense spot of the control was observed.
2.6 RT-PCR Confirmation of Splicing Defects Identified by Microarray
Analysis of prp28∆ YHC1-1 Cells
Strain YTC218 (prp28∆ YHC1-1) was transformed with either plasmid pCA8032 (PRP28/URA3) or an empty pRS316 (URA3) vector and transformants
were grown at 30˚C in SD –Ura liquid medium to an OD600 of 0.5 at which time half of the cells were shifted to 16˚C for 16 hours. The remaining cells were harvested by brief centrifugation, frozen in liquid nitrogen and stored at –80˚C.
After collecting the cells from the 16˚C cultures in the same manner, total RNA was extracted using a hot phenol extraction. Cell pellets were resuspended in
200 µl of TE buffer (10 mM Tris-Cl, 1 mM EDTA) that had been pre-warmed to
70 65˚C and an equal volume of phenol (pH 4.3, Fisher) was added. Cells were incubated at 65˚C for 40 minutes with 30 seconds of vortexing every five minutes. The cell debris and protein was pelleted by centrifugation at 16.1 g for five minutes in a microcentrifuge. The supernatant was transferred to a new tube and extracted two more times at room temperature using phenol/chloroform (pH
5.3, Fisher). Total RNA was then recovered by ethanol precipitation and resuspended in water. Contaminating DNA was removed by RQ1 DNase
(Invitrogen) digestion after which the reaction was again cleaned by a phenol/chloroform (pH 5.3, Fisher) extraction and ethanol precipitation. The resulting total RNA was resuspended in water and used for reverse transcription
(RT). For RT reactions 1 µg of total RNA was annealed with 1 mM exon 2 primer
(see below) in RT reaction buffer supplied by Promega [50 mM Tris-HCl (pH 8.3),
8 mM MgCl2, 50 mM NaCl and 1 mM DTT] by incubating in a Perkin-Elmer thermocycler at 94˚C for 1 minute, 65˚C for 1 minute, 37˚C for 1 minute and then cooling to 4˚C. To the annealed RNA/primer mixtures 1 mM each dNTP, 10 mM
MgCl2, 5 mM DTT and 1 U AMV reverse transcriptase (Promega) were added and the reactions were incubated at 37˚C for 30 minutes, 65˚C for 10 minutes and then cooled to 4˚C. The RT product was then used in subsequent PCR
71 reactions to amplify the spliced and unspliced messages of RPL36B and QCR9.
The U1 snRNA was also amplified as an intronless control. For PCR, 2.5 µl of the RT reaction from above was mixed with 20 mM Tris-HCl(pH 8.4), 50 mM KCl,
2 mM each dNTP, 2 mM MgCl2, 0.25 µM Exon 1 and Intron primers and 0.5 µM
Exon 2 primer, and 2.5 U of platinum Taq polymerase (Invitrogen). The U1 snRNA was amplified using primers U1-2 (5’ gacgttaagcatttatcattgaac 3’) and
U1-3 (5’ cccgttcctaccaagaccttcc 3’). The primers used for amplification of
RPL36B transcripts were RPL36B-EX1 (exon 1 primer, 5’ atggctgtcaagactg 3’),
RPL36B-INT (intron primer, 5’ agtactccgtggagtctttg 3’), and RPL36B-EX2 (exon 2 primer, 5’ taatgacgacgagaggcagc 3’). PCR was conducted by initially heating to
94˚C for 2 minutes followed by 30 cycles of denaturing at 94˚C for 1 minute, annealing at 45˚C for 1 minute and extension at 72˚C for 1 minute. After 30 cycles of PCR, a ten minute 72˚C extension was included and then reactions were cooled to 4˚C. For the amplification of QCR9 products the primers used were QCR9-EX1 (exon 1 primer, 5’ aagcaacaatagcaatacggactaa 3’), QCR9-INT
(intron primer, 5’ tggtgacagcagctctatgaatatg 3’) and QCR9-EX2 (exon 2 primer, 5’ ttactcatcatcgtcgtctccatcg 3’). Reverse transcription and PCR reactions were
72 performed as above except that an annealing temperature of 48˚C was used during PCR. Following PCR, products were visualized by separation on a 1.8% low molecular weight agarose gel.
2.7 Screening for Mutations Synthetically Lethal with prp28∆ YHC1-1
The screen for alleles that are synthetically lethal with prp28∆ YHC1-1 utilized a red/white colony sectoring assay (Koshland et al., 1985) and was conducted in YTC302 [MAT a prp28::HIS3 YHC1-1ade2 ade3 pCA8105
(PRP28/ADE3/URA3)]. Plasmid pCA8105 (PRP28/ADE3/URA3) contains the 3.7 kb SalI-NotI ADE3 fragment from pHT4467 (kindly provided by J. Vanema) in the
SalI-NotI sites of pCA8009 (PRP28/URA3). The sectoring assay is based on the fact that yeast strains lacking the ade2 gene are pink or red due to the accumulation of a pigmented intermediate in the adenine synthesis pathway.
Cells that lack both the ade2 and ade3 genes, however, are white, as mutations in ade3 block synthesis prior to the point of pigment accumulation (Jones, 1981).
This assay therefore allows the detection of colonies that retain a plasmid carrying ADE3 as they are pink or red in color while colonies that have lost the
ADE3 plasmid are white in color. YTC302 cells were grown in YPD (1% yeast
73 extract, 2% bacto-peptone and 2% glucose), diluted and plated to YPD containing 4% glucose (4% YPD). Plates were exposed to UV light to obtain an
80% - 90% killing rate and incubated at 30°C. Colonies that did not appear to sector were picked and re-streaked twice to 4% YPD. Those that remained non- sectoring were tested on media containing 1 mg/mL 5-FOA to select against cells containing the URA3 gene; those that were 5-FOA sensitive were transformed with pCA8034 (PRP28/LEU2). Candidates were then able to sector and became
5-FOA resistant were tested for reversion of the YHC1-1 locus by introducing pYHC1003 (YHC1-1/LEU2) and those which remained 5-FOA sensitive were selected.
2.8 Characterization and Cloning of prp8-501
The one strain that met all of the requirements from the screen is YTC303
[MATαprp28::HIS3 YHC1-1 prp8-501pCA8105 (pRS316-ADE3-PRP28)]. This was crossed to a non-mutagenized isogenic strain YTC301 [MAT a prp28::HIS3
YHC1-1 PRP8 pCA8105 (PRP28/ADE3/URA3)]. Sectoring and viability on 5-
FOA were restored, indicating the mutation is recessive. Tetrad analysis of the resulting diploid yielded 2:2 segregation for survival on 5-FOA indicating that a
74 single gene had been mutated. The gene that had been mutated was identified by complementation with a wild type YCp50-LEU2 library. Transformants that regained the ability to sector were chosen and the library plasmid was recovered and sequenced. The library clone contained only the PRP8 open reading frame, revealing that a mutation in PRP8 was synthetically lethal with prp28∆ YHC1-1.
The library plasmid containing wild type PRP8 is pPRP8001 (YCp50-LEU2-
PRP8). The region of PRP8 containing the synthetic lethal mutation was identified by gap repair using pPRP8002 (PRP8/TRP1) that was gapped separately with three enzymes, AflII, AgeI, and BstEII. Linearized gapped plasmids were then transformed into YTC303 [MAT a prp28::HIS3 YHC1-1 prp8-
501pCA8105 (PRP28/ADE3/URA3)]. The linearized plasmids use the chromosomal copy of PRP8 as a template to repair themselves. Plasmids that copy regions of PRP8 that are still wild-type will now contain a normal copy of the gene, meaning that the yeast colonies will no longer need to retain the PRP28 plasmid. When the region of PRP8 containing the mutation that caused synthetic lethality was copied, no wild-type version of PRP8 would be present and these colonies would still retain the PRP28 plasmid. Only the plasmid that was gapped
75 with BstEII plasmid failed to provide relief from synthetic lethality as judged by sectoring and survival on 5-FOA. This plasmid was rescued and sequenced to identify prp8-501 and was named pPRP8003.
2.9 Testing PRP8 Mutations for Synthetic Lethality with prp28∆ YHC1-1
Mutant alleles of prp8 were obtained from other laboratories (see table
2.2) and tested for synthetic lethality with prp28∆ YHC1-1. The PRP8 alleles obtained from A. Kuhn were carried on pRS313 (CEN/HIS3) and were subsequently sub-cloned into pRS424 (2µ/TRP1) as follows. Plasmids pPRP8026, pPRP8027 and pPRP8028 were created by swapping the BstEII fragment of the pRS313 plasmids carrying alleles prp8-201, prp8-N1809D and prp8-I1851L, respectively into pPRP8025 (PRP8/2µ/TRP1). Plasmid pPRP8025
(PRP8/2µ/TRP1) contains wild-type PRP8 in a XhoI/SacII fragment from pPRP8002 (PRP8/CEN/TRP1). Plasmids pPRP8067, pPRP8068, and pPRP8069 were created by swapping the AgeI fragment of the pRS313 plasmids carrying alleles prp8-R236G, prp8-L280P, and prp8-K611R, respectively into pPRP8025 (PRP8/2µ/TRP1). Plasmids pPRP8070 and pPRP8071 were created by swapping the SexAI/SwaI fragment of pRS313 plasmids carrying alleles prp8-
76 E788G and prp8-D1094A, respectively into pPRP8025 (PRP8/2µ/TRP1). All of the above constructs created by the exchange of PRP8 fragments were confirmed by sequencing.
The plasmids carrying the mutant alleles of PRP8 were tested for synthetic lethality with prp28∆ YHC1-1 by introduction into YTC307 [prp28::HIS3
YHC1-1 prp8::LYS2 pJU169 (PRP8/ URA3)], which was constructed by transforming the 7.5 kb ApaI/SacI fragment containing prp8∆::LYS2 from pJU224
(prp8∆::LYS2/TRP1) into YTC300 (prp28::HIS3 YHC1-1) containing pJU169
(PRP8/ URA3). Lys+ transformants were identified and homologous recombination of prp8∆::LYS2 at the correct chromosomal locus was confirmed by southern blot (data not shown). Transformation of prp8 mutant alleles was followed by selection on media containing 1 mg/mL 5-FOA. Plasmids that conferred synthetic lethality were rescued from the yeast strain and subjected to sequencing to verify the mutation within PRP8 that caused lethality. To assure that the PRP8 alleles causing cell inviability on 5-FOA were synthetically lethal and not null mutations, plasmids were transformed into YJU75 [prp8::LYS2 pJU169 (PRP8/ URA3)] and plated on media containing 1 mg/mL 5-FOA.
77 2.10 Back-crossing of Bypass Suppressor Strains
Two bypass candidate strains (MAT a prp28::TRP1 yhc1::HIS3 xbp pYHC1005 (YHC1/LEU2)] were back-crossed to strain YTC286 [MAT α prp28::TRP1 pCA8032 (PRP28/URA3)] three times in order to remove unwanted background mutations introduced by UV mutagenesis. Strains were patched together on YPD and then streaked to selective SD medium to isolate diploids.
The diploids were then patched to YPD before being transferred to 1.5 % potassium acetate solution and shaken at 30˚C for 2-7 days until spores were visible. Spores were collected by centrifugation in a microcentrifuge at 0.8 g for 1 minute and washed twice with water. 20 µl of washed spores were mixed with 5
µl of ß-glucuronidase (SIGMA) and incubated at 37˚C for 5 minutes to digest the ascus prior to tetrad dissection on YPD. After tetrads were grown, their genotypes were tested by replica plating to selective SD medium to isolate the desired strains.
78 2.11 Identification of prp42-101 and snu71-101 as a Bypass Suppressors of
PRP28
To identify the genes containing the bypass mutations in the two strains recovered from the genetic screen, a wild-type yeast genomic library (YCp50-
LEU2) was transformed into strains isolated after the first round of back-crossing
[MAT a prp28::TRP1 xbp pCA8032 (PRP28/URA3)]. Transformants were replica- plated to SD-Leu plates containing 1 mg/mL 5-FOA. Colonies that were 5-FOA sensitive were selected and re-tested on 5-FOA medium to confirm their dependence on pCA8032 (PRP28/URA3). Candidate strains were subjected to plasmid rescue to recover the plasmids from the cell. As these strains contained both the desired library plasmid as well as pCA8032 (PRP28/URA3), recovered plasmids were digested with HindIII (New England Biolabs) to determine which plasmid had been recovered. For the first candidate, eight of the thirteen rescued plasmids were pCA8032 while the remaining five had the same digestion pattern that was distinct from that of pCA8032. For the second candidate, six of the ten rescued plasmids were pCA8032. The remaining four demonstrated the same novel digestion pattern that was distinct from pCA8032 as well as the library plasmid identified from the first candidate. Re-
79 transformation of these plasmids into their respective bypass strains conferred sensitivity to 5-FOA, thus confirming the presence of the wild-type gene of interest on these plasmids. The plasmids were then subjected to sequencing utilizing primers YCp50-1 (5’ ccatacccacgccg-aaacaagcgctc 3’) and YCp50-2 (5’ atatgcgttgatgcaatttctatgcgcac 3’). These primers anneal to the YCp50 vector adjacent to the library DNA insertion site, thus allowing the sequencing of the ends of the library insert. The resulting sequences were then used for a BLAST search of the S. cerevisiae genome database (Dolinski, 2001) in order to identify the open reading frames included in the library insert. The library plasmid recovered from the first bypass candidate strain contained a 12-kilobase (kb) library insert that mapped to chromosome IV and contained several open reading frames including PRP42. The library plasmid recovered from the second bypass candidate strain contained a 9-kb insert that mapped to chromosome VII and contained several open reading frames including SNU71.
2.12 Cloning of Wild-type and Mutant PRP42
The wild-type version of PRP42 was sub-cloned from the wild-type library plasmid pPRP42001. Plasmid pPRP42001 was cut with restriction enzymes
80 FspI and SapI and treated with Klenow to create blunt ends (the restriction digest was supplemented with 33 µM each dNTP and 1 µl Klenow fragment and incubated at 25˚C for 15 minutes). The fragment containing PRP42 was isolated and cloned into the SmaI site of pRS315 in the forward orientation to create plasmid pPRP42014. The mutant version of PRP42 was obtained by the
Polymerase Chain Reaction (PCR) using primers p42-3 (5’ ccccggatcctctacc- ttggaaatccca 3’, BamHI site is underlined) and p42-2 (5’ cccccgtcgaccaatgcc- ttttggctaagg 3’, SalI site is underlined). PCR was conducted using genomic DNA
(gDNA) obtained from the bypass suppressor strain as a template. The 100 µl reaction contained 5 µl gDNA, 10 µl 10 x PCR buffer supplied by Invitrogen (500
mM KCl, 100 mM Tris-HCl (pH 8.3), 5 mM MnCl2, 5 µl 50 mM MgCl2, 2 µM 10mM dNTPs (10 mM each dATP, dCTP, dGTP and dTTP), 0.5 µl each 100 µM primer, and 0.7 µl Taq polymerase (Invitrogen). The mutant PCR product was subjected to sequencing as well as being digested with BamHI and SalI and cloned into the same sites of the yeast expression vectors pRS315 and pRS316 to create plasmids pPRP42007 and pPRP42005 respectively.
81 2.13 Cloning of Wild-type and Mutant SNU71
A wild-type allele of SNU71 was obtained from pSNU71001, the original wild-type library plasmid isolated (SNU71/LEU2). A SmaI/NdeI fragment that contained SNU71 was removed from pSNU71001, treated with Klenow to create blunt ends (see above for details of Klenow treatment), and inserted into the
SmaI site of pRS315 to create plasmid pSNU71011 (SNU71/LEU2). The
BamHI/SalI fragment from pSNU71011 was then inserted into the same sites of pRS316 to create pSNU71016 (SNU71/URA3). The mutant allele snu71-101 was obtained by PCR (see above) using gDNA from the candidate strain and primers snu71-1 (5’ cccccggatccgacactatcgtagaaggg 3’, BamHI site is underlined) and snu71-2 (5’ cccccgtcgacctgtttcagagcgagcctt 3’, SalI site is underlined). The PCR product was submitted for sequencing as well as being used for plasmid construction. The PCR fragment was initially cut with BamHI and SalI and inserted into the same sites of pRS315 to create plasmid pSNU71003. After finding that this plasmid was not functional due to an incomplete 3’ untranslated region (UTR), an SphI/BssHII fragment containing the snu71-101 mutation was moved into pSNU71011 (SNU71/LEU2) to create
82 plasmid pSNU71015 (snu71-101/LEU2). The entire gene was then moved into pRS314 using the BamHI and SalI sites to make pSNU71017 (snu71-101/TRP).
2.14 Analyzing the Growth Phenotype of prp28∆ prp42-101
YTC727 (MAT a prp28::TRP1 prp42-101) was transformed with either pCA8032 (PRP28/URA3) or pRS316 (URA3). These transformants as well as
YTC286 [MAT α prp28::TRP1 pCA8032 (PRP28/URA3)] were grown in SD-Ura
liquid medium to saturation. Cells corresponding to 0.5 OD600 units were collected and resuspended in 100µl of water. Three ten-fold serial dilutions were performed and 4 µl of each dilution were spotted to four duplicate SD-Ura plates.
Plates were incubated at 16˚C, 25˚C, 30˚C and 37˚C until growth in all spots of the control strain was apparent.
2.15 prp42-101 Does not Produce a Growth Defect on its Own
Strain YTC634 (MAT a/α prp42::KAN/+) was purchased from ResGen
(ResGen strain YDR235W), transformed with plasmid pPRP42015
(PRP42/URA3), subjected to sporulation and tetrad dissection in order to isolate haploid deletion strain YTC635 [MAT a prp42::KAN pPRP42004 (PRP42/URA3)].
83 YTC635 was subsequently transformed with plasmids pPRP42014
(PRP42/LEU2) and pPRP42007 (prp42-101/LEU2). Resulting transformants were streaked to SD-Leu plates containing 1 mg/mL 5-FOA to select cells that did not contain pPRP42015 (PRP42/URA). Colonies from the 5-FOA plate were inoculated into SD-Leu liquid medium and grown to saturation. Cells
corresponding to 0.5 OD600 units were collected and resuspended in 100µl of water. Three ten-fold serial dilutions were performed and 4 µl of each dilution were spotted to four duplicate SD-Ura plates. Plates were incubated at 16˚C,
25˚C, 30˚C and 37˚C until growth in all spots of the control strain was apparent.
2.16 Deletion of PRP42 Cannot Bypass PRP28
Strain YTC635 [MAT a prp42::KAN pPRP42004 (PRP42/URA3)] was transformed with pPRP42007 (prp42-101/LEU2) and a transformant was streaked to plates containing 1 mg/mL 5-FOA to select cells which had lost pPRP42004 (PRP42/URA3), resulting in a prp42::KAN prp42-101/LEU2 strain.
This strain was subsequently transformed with a prp28::HIS3 fragment [2.3-kb
EcoRI/NotI fragment from plasmid pCA8033 (prp28::HIS3/KS+)]. His+ colonies were replica plated and incubated at 16˚C to identify colonies that were cold-
84 sensitive. Genomic DNA from the candidates was isolated and then used for a
PCR reaction to determine whether recombination of the prp28::HIS3 fragment occurred at the correct locus. PCR was carried out using primers 28-27 (5’ ctgagcaacttcctacgtccatgtg 3’), HIS3D (5’ cagtggtgtgatggtcgtctatgtg 3’) and
PRP28-TAP2 (5’ tcccggaatcagaaaggcctgatgt 3’) to detect both wild-type and deletion versions of PRP28. Correct candidates were further confirmed by
Southern analysis using the HindIII/BglII fragment of pCA8033 (prp28::HIS3/KS+) as probe to gDNA digested with HindIII. A correct strain, YTC799 [MAT a prp28::HIS3 prp42::KAN pPRP42007 (prp42-101/LEU2)] was transformed with pPRP42017 (PRP28/PRP42/URA3). After transformation a colony which had lost pPRP42007 (prp42-101/LEU2) was selected. These cells were then streaked to SD complete media containing 1 mg/mL 5-FOA and incubated at
30˚C until growth of single colonies in the control was apparent.
2.17 prp42-101 and snu71-101 are Synthetically Lethal with YHC1 Bypass
Alleles
To construct a strain to test the synthetic lethality of prp42-101 with the alleles of YHC1, YTC513 [MAT α yhc1::HIS3 pYHC1023 (YHC1-HA/URA3)] was
85 trasformed with pYHC1005 (YHC1/LEU2) and streaked to SD-Leu medium containing 1 mg/mL 5-FOA to select cells without pYHC1023 (YHC1-HA/URA3).
The resulting cells were transformed with pPRP42009 (PRP42/YHC1/URA3) and transformants were replica-plated to SD-Leu plates to identify colonies that had lost pYHC1005 (YHC1/LEU2). This process resulted in a strain that was MAT α yhc1::HIS3 pPRP42009 (PRP42/YHC1/URA3). Simultaneously, strain YTC635
[MAT a prp42::KAN pPRP42004 (PRP42/URA3)] was transformed with plasmid pPRP42010 (prp42-101/LYS2). Transformants were streaked to media containing 1 mg/mL 5-FOA to counter-select plasmid pPRP42004
(PRP42/URA3). This strain [MAT a prp42::KAN pPRP42010 (prp42-101/LYS2)] was then mated to the above strain [MAT α yhc1::HIS3 pPRP42009
(PRP42/YHC1/URA3)]. After mating diploids were selected, sporulated and tetrads were dissected to isolate strain YTC762 [MAT α yhc1::HIS3 prp42::KAN pPRP42009 (YHC1/PRP42/URA3)]. This strain was re-transformed with pPRP42010 (prp42-101/LYS2), as it was not maintained after tetrad dissections.
Subsequently this strain was transformed with the pool of YHC1 L13 mutant alleles carried on LEU2 marked plasmids (see table 2.2). These transformants were grown in either SD-Lys-Leu or SD-Ura-Lys-Leu liquid medium and cells
86 corresponding to 0.5 OD600 units were collected and resuspended in 100µl of water. Three ten-fold serial dilutions were performed and 4 µl of each dilution were spotted to SD-Lys-Leu plates containing 5-FOA. Plates were incubated at
30˚C, 25˚C and 16˚C until growth was seen in the lowest dilution of the control.
To construct a strain for testing the synthetic lethality of snu71-101 with the YHC1 alleles YTC764 [MAT α snu71::KAN pSNU71001 (SNU71/LEU2)] was first crossed to a wild-type strain with the W303 background YTC442 (MAT a ade2 ura3 trp1 his3 leu2 can-100) in order to provide a healthier strain with good sporulation. The resulting strain was transformed with pSNU71016
(SNU71/URA3) and a colony that had lost pSNU71001 (SNU71/LEU2) was identified by replica-plating, yielding strain YTC783 [MAT α snu71::KAN pSNU71016 (SNU71/URA3)]. This strain was subsequently transformed with pSNU71017 (snu71-101/TRP) and streaked to plates containing 1 mg/mL 5-FOA to select cells that had lost pSNU71016 (SNU71/URA3). The resulting strain is
[MAT α snu71::KAN pSNU71017 (snu71-101/TRP)]. Simultaneously, strain
YTC511 [MAT a yhc1::HIS3 pYHC1023 (YHC1-HA/2µ/URA3)] was transformed with plasmid pYHC1005 (YHC1/LEU2) followed by selection on plates containing
1 mg/mL 5-FOA to identify cells that had lost pYHC1023 (YHC1-HA/2µ/URA3).
87 This strain was then transformed with plasmid pSNU71018
(SNU71/YHC1/URA3) which was constructed by ligating a BamHI fragment containing YHC1 from pYHC1005 (YHC1/LEU2) into pSNU71016
(SNU71/URA3). Transformants were replica-plated to identify colonies that had lost pYHC1005 (YHC1/LEU2). This resulting strain [MAT a yhc1::HIS3 pSNU71018 (SNU71/YHC1/URA3)] was mated to the above strain [MAT α snu71::KAN pSNU71017 (snu71-101/TRP)]. Diploids were selected and then subjected to sporulation and tetrad dissection to obtain strain YTC793 [MAT a yhc1::HIS3 snu71::KAN pSNU71018 (SNU71/YHC1/URA3)]. Plasmids containing the YHC1 L13 alleles (pRS315) were transformed into this strain and transformants were grown in SD-Lys-Leu liquid medium. Cells corresponding to
0.5 OD600 units were collected and resuspended in 100 µl of water. Three ten- fold serial dilutions were performed and 4 µl of each dilution were spotted to SD-
Lys-Leu plates containing1 mg/mL 5-FOA. Plates were incubated at 30˚C, 25˚C and 16˚C until growth was seen in the lowest dilution of the control.
88 2.18 Alleles of prp8 are Synthetically Lethal with prp42-101
Strain YTC754 [MAT a prp42::KAN pPRP42007 (prp42-101/LEU2)] was crossed to strain YTC786 [MAT α prp8::LYS2 pPRP8025 (PRP8/URA3)].
Diploids were isolated and subjected to sporulation and tetrad dissection to obtain strain YTC835 [MAT a prp42::KAN prp8::LYS2 pPRP42007 (prp42-
101/LEU2) pPRP8025 (PRP8/URA3)]. YTC835 was transformed with the 2.3 kb
EcoRI/NotI prp28::HIS3 fragment from plasmid pCA8033 (prp28::HIS3/KS+).
Recombination at correct locus was confirmed by PCR to detect both wild-type and deletion versions of PRP28 using primers 28-27 (5’ ctgagcaacttcc- tacgtccatgtg 3’), HIS3D (5’ cagtggtgtgatggtcgtctatgtg 3’) and PRP28-TAP2 (5’ tcccggaatcagaaaggcctgatgt 3’). The resulting strain is YTC836 [MAT a prp28::HIS3 prp42::KAN prp8::LYS2 pPRP42007 (prp42-101/LEU2) pPRP8025
(PRP8/URA3)] which was then transformed with each of the mutant alleles of prp8 carried on TRP1 marked plasmids (see table 2.2). Transformants were selected on SD-Trp and then streaked to SD-Trp for purification. Single colonies were then inoculated into SD-Trp media and grown to saturation. Cells
corresponding to 0.5 OD600 units were collected and resuspended in 100µl of water. Three ten-fold serial dilutions were performed and 4 µl of each dilution
89 were spotted to SD plates containing 5-FOA. Plates were incubated at 30˚C for approximately three days until single colonies appeared in the least concentrated spot of the positive control.
2.19 Testing Tail-truncated Sm Proteins for bypass of Prp28p
Strains containing Sm tail truncations were obtained from M. Rosbash and include DZY6 (YTC621) [smb::KAN pDZ1=pSM007(SmB∆C/TRP1)], DZY19
(YTC623) [smd1::LEU2 pDZ7=pSM008 (SmD1∆C/TRP1)], DZY20 (YTC624)
[smd3∆1::LEU2 pDZ8=pSM009(SmD3∆C/TRP1)] and DZY21 (YTC625)
[smb::KAN smd3∆1::LEU2 pDZ3=pSM011 (SmB∆C/SmD3∆C/TRP1)] (Zhang et al., 2001). These strains were crossed to prp28∆ strains of the opposite mating type, either strain YTC62 (MAT a) or YTC65 (MAT α) [prp28::HIS3 pCA8032
(PRP28/URA3)], to obtain strains YTC626 [MAT a smb::KAN prp28::HIS3 pDZ1
(SmB∆C/TRP1) pCA8032 (PRP28/URA3)], YTC628 [MAT a smd1::LEU2 prp28::HIS3 pDZ7 (SmD1∆C/ TRP1) pCA8032 (PRP28/URA3)], YTC630 [MAT α smd3::LEU2 prp28::HIS3 pDZ8 (SmD3∆C/ TRP1) pCA8032(PRP28/URA3)], and
YTC632 [MAT α smb::KAN smd3::LEU prp28::HIS3 pDZ3 (SmB∆C/SmD3∆C/
90 TRP1) pCA8032 (PRP28/URA3)]. These strains were then tested on YP plates containing 3% galactose, 1% sucrose and 1 mg/mL 5-FOA.
2.20 Deletion of MUD1 Cannot Bypass PRP28
Strain YTC765 (MAT a mud1::KAN) (ResGen strain YBR119W) was mated to YTC65 [MAT α prp28::HIS3 pCA8032 (PRP28/URA3)]. Diploids were subjected to sporulation and tetrad dissection. Tetrads were replica-plated to selective media revealing that spores which were [mud1:KAN prp28::HIS3 pCA8032 (PRP28/URA3)] were unable to survive in the presence of 5-FOA.
These spores were subsequently streaked to plates containing 5-FOA to confirm this finding.
91
Strain Genotype Source YTC62 MAT a prp28::HIS3 ura3-52 lys2-801 ade2-101 Chang Lab trp1-∆1 his3∆-200 leu2-∆1 (pCA8032)
YTC65 MAT α prp28::HIS3 ura3-52 lys2-801 ade2-101 Chang Lab trp1-∆1 his3∆200 leu2-∆1 (pCA8032)
YTC102 MAT α prp28::HIS3 ura3-52 lys2-801 ade2-101 Chang Lab trp1-∆1 his3∆200 leu2-∆1 (pCA8070)
YTC105 MAT a ura3-52 lys2-801 ade2-101 trp1-∆1 Chang Lab his3-∆200 leu2-∆1
YTC218 MAT α prp28::HIS3 YHC1-1 ura3-52 lys2-801 Chang Lab ade2-101 trp1-∆1 his3∆200 leu2-∆1
YTC286 MAT α prp28::TRP1 ura3-1 ade2-1 trp1-1 Chang Lab his3-11 leu2-3,112 can1-100 (pCA8032)
YTC300 MAT a prp28∆ YHC1-1 ura3-52 lys2-801 ade2-101 This study ade3 trp1-∆1 his3∆200 leu2-∆1
YTC301 MAT α prp28::HIS3 YHC1-1 ura3-52 lys2-801 This study ade2-101 ade3 trp1-∆1 his3∆200 leu2-∆1 (pCA8105)
YTC302 MAT a prp28::HIS3 YHC1-1 ura3-52 lys2-801 This study ade2-101 ade3 trp1-∆1 his3∆200 leu2-∆1 (pCA8105)
YTC303 MAT a prp28::HIS3 YHC1-1 prp8-501 ura3-52 This study lys2-801ade2-101 ade3 trp1-∆1 his3∆200 leu2-∆1 (pCA8105)
YTC304 MAT a prp8::LYS2 cup1∆::ura3 ura3 lys2 ade2 C. Guthrie his3 leu2 (pJU169) (continued)
Table 2.1 Yeast Strains
92 Table 2.1 (continued)
Strain Genotype Source YTC307 MAT α prp28::HIS3 YHC1-1 prp8::LYS2 ura3-52 This study lys2-801ade2-101 ade3 trp1-∆1 his3∆200 leu2-∆1 (pJU169)
YTC442 MAT a ura3-1 ade2-1 trp1-1 his3-11 leu2-3,112 Chang Lab can1-100
YTC510 MAT a yhc1::HIS3 ura3-1 trp1-1 his3-11 leu2-3,112 Chang Lab can1-100
YTC513 MAT α yhc1::HIS3 ura3-1 lys2∆::hisG ade2-1 Chang Lab trp1-1 his3-11 leu2-3,112 can1-100 (pYHC1023)
YTC621 MAT a smb::KAN ura3-52 trp1-289 his3-∆1 M. Rosbash (DZY6) leu2-3, -112 arg4 (pSM007) (pDZ1)
YTC623 MAT α smd1::LEU2 ura3-52 trp1-289 his3-∆1 M. Rosbash (DZY19) leu2-3, -112 arg4 (pSM008) (pDZ7)
YTC624 MAT α smd3∆1::LEU2 ura3-52 trp1-∆101 his1 M. Rosbash (DZY20) leu2-1 (pSM009) (pDZ8)
YTC625 MAT α smb::KAN, smd3∆1::LEU2 trp1-∆101 M. Rosbash (DZY21) ura3-52 leu2-3, -112 (pSM011) (pDZ3)
YTC626 MAT a smb::KAN prp28::HIS3 trp1 leu2 ura3-52 This study his3 ade2 (pSM007) (pDZ1) (pCA8032)
YTC628 MAT a smd1::LEU2 prp28::HIS3 trp1 ura3-52 his3 This study leu2 ade2 (pSM008) (pDZ7) (pCA8032)
YTC630 MAT α smd3::LEU2 prp28::HIS3 leu2 ura3-52 This study trp1 (pSM009) (pDZ8) pCA8032 (continued)
93 Table 2.1 (continued)
Strain Genotype Source YTC632 MAT α smb::KAN smd3::LEU prp28::HIS3 trp1 This study ura3-52 leu2 ade2 (pSM011) (pDZ3) (pCA8032)
YTC634 MAT a/α prp42::KAN/+ ura3∆0/ura3D0 lys2∆0/+ ResGen his3∆0/his3∆0 leu2∆0/leu2∆0 met15∆o/+
YTC635 MAT a prp42::KAN ura3∆0 lys2D0 his3∆0 leu2∆0 This study (pPRP42004)
YTC727 MAT a prp28::TRP prp42-101 ura3-1 lys2 ade2-1 This study trp1-1 his3-11 leu2-3,112 can1-100
YTC728 MAT a prp28::TRP prp42-101 ura3-1 lys2 ade2-1 This study trp1-1 his3-11 leu2-3,112 can1-100 (pCA8032)
YTC729 MAT α prp28::TRP prp42-101 ura3-1 lys2 ade2-1 This study trp1-1 his3-11 leu2-3,112 can1-100
YTC730 MAT α prp28::TRP prp42-101 ura3-1 lys2 ade2-1 This study trp1-1 his3-11 leu2-3,112 can1-100 (pCA8032)
YTC731 MAT a prp28::TRP snu71-101 ura3-1 lys2 ade2-1 This study trp1-1 his3-11 leu2-3,112 can1-100
YTC732 MAT a prp28::TRP snu71-101 ura3-1 lys2 ade2-1 This study trp1-1 his3-11 leu2-3,112 can1-100 (pCA8032)
YTC733 MAT α prp28::TRP snu71-101 ura3-1 lys2 ade2-1 This study trp1-1 his3-11 leu2-3,112 can1-100
YTC734 MAT α prp28::TRP snu71-101 ura3-1 lys2 ade2-1 This study trp1-1 his3-11 leu2-3,112 can1-100 (pCA8032)
YTC754 MAT a prp42::KAN cup1::ura3 ura3 lys2 trp1 his3 This study leu2 (pPRP42010) (continued)
94 Table 2.1 (continued)
Strain Genotype Source YTC762 MAT α yhc1::HIS3 prp42::KAN ura3 lys2 ade2-1 This study trp1-1 his3 leu2 (pPRP42017)
YTC764 MAT α snu71::KAN ura3∆0 lys2D0 his3∆0 leu2∆0 ResGen (pSNU71001)
YTC765 MAT a mud1::KAN ura3 his3 leu2 met ResGen
YTC783 MAT α snu71::KAN ura3 lys2 his3 leu2 This study (pSNU71001)
YTC786 MAT α prp8::LYS2 ura3 ade2 lys2 trp1 his3 leu2 C. Guthrie (pJU169)
YTC793 MAT a yhc1::HIS3 snu71::KAN ura3 lys2 ade2 This study trp1-1 leu2 (pSNU71017) (pSNU71019)
YTC799 MAT a prp28::HIS1 prp42::KAN ura3∆0 lys2∆0 This study his3∆0 leu2∆0 (pPRP42007)
YTC815 MAT a prp28::TRP1 yhc1::HIS3 ura3 lys2 ade2 This study trp1 his3 leu2 can1-100 (pYHC1054)
YTC835 MAT a prp42::KAN prp8::LYS ura3 lys2 ade2 This study trp1 his3 leu2 (pPRP42007) (pPRP8025)
YTC836 MAT a prp28::HIS3 prp42::KAN prp8::LYS This study ura3 lys2 ade2 trp1 his3 leu2 (pPRP42007) (pPRP8025)
95
Plasmid Description Source pCA8005 PRP28 in KS (+). A ~3.6 Kb KpnI fragment Chang lab containing the entire PRP28 gene was cloned into KS(+)/KpnI. pCA8009 PRP28 in pRS316. A ~3.6 Kb KpnI fragment Chang lab containing the entire PRP28 gene was cloned into pRS316/KpnI. pCA8032 PRP28 in pRS316. A KpnI-BclI fragment from Chang lab pCA8005 was cloned into pRS316/KpnI/BamHI. pCA8033 prp28::HIS3 in KS (+). The internal BglII Chang lab fragment of PRP28 was removed and replaced with a BamHI fragment containing the HIS3 gene. pCA8065 pRSET-A-PRP28. A 2.4-Kb BamHI fragment Chang lab ([IEGR/Xa protease site]-ATG-PRP28-EcoRI-20 bp) was subcloned into pRSET-A for overexpressing
(His)6-Prp28p. pHT4467 ADE3/URA3. ADE3 in the SmaI/SpeI sites J. Venema of pRS316 pPRP8001 PRP8 in YCp50/LEU2. Original library plasmid This study isolated during the prp28∆ YHC1-1 synthetic lethal screen. pPRP8002 PRP8 in pRS314. The BsiEI/BsrBI fragment from This study pPRP8001 was inserted into the SmaI site of pRS314. pPRP8003 prp8-501 in pRS314. pPRP8002 was gapped This study with BstEII and transformed into the synthetic lethal strain for gap repair. This plasmid was rescued and sequenced. (continued) Table 2.2 Plasmids
96 Table 2.2 (continued)
Plasmid Description Source pPRP8004 prp8::LYS2 in pRS424. Plasmid pJU224 C. Guthrie constructed by Jim Umen. pPRP8006 prp8-101 (E1960K) in pRS424. Plasmid C. Guthrie pJU232 constructed by Jim Umen. pPRP8007 prp8-102 (E1960G) in pRS424. Plasmid C. Guthrie pJU228 constructed by Jim Umen. pPRP8008 prp8-103 (F1834L) in pRS424. Plasmid C. Guthrie pJU231 constructed by Jim Umen. pPRP8009 prp8-104 (F1834S) in pRS424. Plasmid C. Guthrie pJU230 constructed by Jim Umen. pPRP8010 prp8-121 (W1608R) in pRS424. Plasmid C. Guthrie pJU254 constructed by Jim Umen. pPRP8011 prp8-122 (W1574R) in pRS424. Plasmid C. Guthrie pJU255 constructed by Jim Umen. pPRP8012 prp8-123 (E1575V) in pRS424. Plasmid C. Guthrie pJU256 constructed by Jim Umen. pPRP8013 prp8-124 (M1397I) in pRS424. Plasmid C. Guthrie pJU256 constructed by Jim Umen. pPRP8014 prp8-135 (E1817G) in pRS424. Plasmid C. Guthrie pCC81 constructed by Cathy Collins. pPRP8015 prp8-136 (N1869D) in pRS424. Plasmid C. Guthrie pCC82 constructed by Cathy Collins. pPRP8016 prp8-143 (K1864E) in pRS424. Plasmid C. Guthrie pCC86 constructed by Cathy Collins.
(continued)
97 Table 2.2 (continued)
Plasmid Description Source pPRP8017 prp8-144 in pRS424. Plasmid pCC85 C. Guthrie constructed by Cathy Collins. pPRP8018 PRP8/URA3/CEN. Plasmid constructed C. Guthrie by Jim Umen. pPRP8019 prp8-125 (T1564A) in pRS424. Plasmid C. Guthrie pJU258 constructed by Jim Umen. pPRP8020 prp8-501 in pRS424. A SacII/XhoI fragment This study was removed from pPRP8003 and inserted into the SacII/XhoI sites of pPRP8005. Presence of mutations confirmed by sequencing pPRP8021 prp8-501 mutations in pRS314. pPRP8003 This study and pPRP8002 were digested with BstEII, the prp8-501 insert from pPRP803 was inserted into pPRP8002. Presence of the mutations was confirmed by sequencing. pPRP8022 prp8-201 (T1861P) in pRS313 A. Kuhn pPRP8023 prp8-N1809D in pRS313. A. Kuhn pPRP8024 prp8-I1851L in pRS313. A. Kuhn pPRP8025 Wild type PRP8 in pRS424. The XhoI/SacII This study fragment of pPRP8002 was ligated into the same sites ofpRS424. pPRP8026 prp8-201 in pRS424. The BstEII fragment from This study pPRP8022 was swapped into pPRP8025. pPRP8027 prp8-N1809D in pRS424. The BstEII fragment This study from pPRP8023 was swapped into pPRP8025.
(continued)
98 Table 2.2 (continued)
Plasmid Description Source pPRP8028 prp8-I1851L in pRS424. The BstEII fragment This study from pPRP8024 was swapped into pPRP8025. pPRP8029 Clone M8 = T1702I in pRS424 M. Konarska pPRP8030 Clone M18 = M1783K in pRS424 M. Konarska pPRP8031 Clone M29 = I1933F in pRS424 M. Konarska pPRP8032 Clone M30 = L1794F in pRS424 M. Konarska pPRP8033 Clone M50 = S1906G in pRS424 M. Konarska pPRP8034 Clone M54 = D1670N, D1778G in pRS424 M. Konarska pPRP8035 Clone B4 = D1700G in pRS424 M. Konarska pPRP8036 Clone B15 = F1883D in pRS424 M. Konarska pPRP8038 Clone B20 = S1769S, D1778G in pRS424 M. Konarska pPRP8039 Clone B30 = I1775T, D1778E in pRS424 M. Konarska pPRP8040 Clone B69 = D 1778G, N1869S in pRS424 M. Konarska pPRP8041 Clone B72 = S1745P, D1778G in pRS424 M. Konarska pPRP8042 Clone B93 = D1778R, L1779M, and G1635D M. Konarska in pRS424 pPRP8043 Clone C13 = S1829C, E1832V in pRS424 M. Konarska pPRP8044 Clone C41 = E1832A, L1891P in pRS424 M. Konarska pPRP8045 Clone C69 = E1832V, P1952L in pRS424 M. Konarska pPRP8046 Clone C96 = K1807R, S1829C, and E1832A M. Konarska in pRS424 pPRP8047 Clone D23 = Q1677R, D1700E in pRS424 M. Konarska
(continued)
99 Table 2.2 (continued)
Plasmid Description Source pPRP8048 Clone E2 = S1745F in pRS424 M. Konarska pPRP8049 Clone G10 = F1724L in pRS424 M. Konarska pPRP8050 Clone H3 = E1842N in pRS424 M. Konarska pPRP8051 CloneH5 = N1792S and E1842N in pRS424 M. Konarska pPRP8052 Clone H9 = Y1787C and E1842N in pRS424 M. Konarska pPRP8054 Clone 8-152 = N1869D and S1970R in pRS424 M. Konarska pPRP8055 Clone 8-152b = S1970R in pRS424 M. Konarska pPRP8056 Clone 8-153 = T1982A in pRS424 M. Konarska pPRP8057 Clone 8-154 = T1982A and SA1966/7AG in M. Konarska pRS424 pPRP8058 Clone 8-155 = T1982A and V1987A in pRS424 M. Konarska pPRP8059 Clone 8-156 = N1869D and T1982A in pRS424 M. Konarska pPRP8060 Clone 8-157 = N1869D, S1970R, T1982A and M. Konarska V1987A in pRS424 pPRP8061 Clone 8-158 = yeast amino acid sequence 1962– M. Konarska 1975 converted to human amino acid sequence 1890-1903 in pRS424 pPRP8062 prp8-R236G in pRS313 A. Kuhn pPRP8063 prp8-L280P in pRS313 A. Kuhn pPRP8064 prp8-K611R in pRS313 A. Kuhn pPRP8065 prp8-E788G in pRS313 A. Kuhn pPRP8066 prp8-D1094A in pRS313 A. Kuhn
(continued)
100 Table 2.2 (continued)
Plasmid Description Source pPRP8067 prp8-R236G in pRS424. The AgeI fragment This study from pPRP8062 was swapped into pPRP8025. pPRP8068 prp8-L280P in pRS424. The AgeI fragment This study from pPRP8063 was swapped into pPRP8025. pPRP8069 prp8-K611R in pRS424. The AgeI fragment This study from pPRP8064 was swapped into pPRP8025. pPRP8070 prp8-E788G in pRS424. The SexAI/SwaI This study fragment from pPRP8065 was swapped into pPRP8025. pPRP8071 prp8-D1094A in pRS424. The SexAI/SwaI This study fragment from pPRP8066 was swapped into pPRP8025. pPRP42001 Wild-type PRP42 in YCp50-LEU2 library vector. This study PRP42 is part of an ~12-kb insert that contains several other ORFs; isolated as library plasmid that suppressed bypass ability of candidate. pPRP42004 Wild-type PRP42 in pRS316. PCR product using This study primers p42-3 and p42-2 was cut with BamHI and SalI and cloned into the same sites of pRS316. pPRP42005 Mutant prp42-101 in pRS316. PCR product using This study primers p42-3 and p42-2 was cut with BamHI and SalI and placed into the same sites of pRS316. pPRP42007 Mutant prp42-101 in pRS315. PCR product using This study primers p42-3 and p42-2 was cut with BamHI and SalI and cloned into the same sites of pRS315.
(continued)
101 Table 2.2 (continued)
Plasmid Description Source pPRP42009 Wild-type PRP42 and wild-type YHC1 in pRS316. This study A BamHI fragment containing YCH1 from pYHC1005 was cloned into the BamHI site of pPRP42004 in the forward orientation.
pPRP42010 Mutant prp42-101 in pRS317. pRS317 and This study pPRP42005 (prp42-101/pRS316) were cut with SalI and NotI, the pPRP42005 insert was put into pRS317.
pPRP42014 Wild-type PRP42 cloned from original library This study plasmid into pRS315. pPRP42001 was cut with FspI and SapI and cloned into the SmaI site of pRS315.
pPRP42015 Wild-type PRP42 in pRS316. pPRP42001 was This study cut with FspI and SapI and cloned into the SmaI site of pRS316.
pPRP42017 PRP28/PRP42 in pRS316. pPRP42014 This study (PRP42/pRS315) and pCA8009 (PRP28/pRS316) were cut with SalI and NotI, the PRP42 fragment was inserted into pCA8009.
pSM007 SmB∆C = nucleotides encoding amino acids M. Rosbash (pDZ1) 1-106 of SmB cloned into BamHI/SalI of pESC-TRP (Stratagene)
pSM008 SmD1∆C = nucleotides encoding amino acids M. Rosbash (pDZ7) 1-125 of SmD1 cloned into EcoRI/ClaI of pESC-TRP (Stratagene)
pSM009 SmD3∆C = nucleotides encoding amino acids M. Rosbash (pDZ8) 1-85 of SmD3 cloned into EcoRI/ClaI of pESC-TRP (Stratagene)
(continued) 102 Table 2.2 (continued)
Plasmid Description Source pSM011 SmB∆C/SmD3∆C = nucleotides encoding amino M. Rosbash (pDZ3) acids 1-106 of SmB and 1-85 of SmD3 cloned into BamHI/SalI and EcoRI/ClaI of pESC-TRP (Stratagene) respectively
pSNU71001 Wild-type SNU71 in YCp50/LEU2 library plasmid, This study rescued from bypass suppressor candidate.
pSNU71003 Mutant snu71-101 in pRS315, incomplete 3’ end. This study PCR fragment from gDNA using primers SNU71-1 and SNU71-2 was cut with BamHI/SalI and inserted into the same sites of pRS315. Although this plasmid contains the snu71-101 mutation, it lacks the entire 3’ UTR and was therefore used only as a source of the snu71-101 mutation for subsequent cloning.
pSNU71011 Wild-type SNU71 in pRS315. The SmaI/NdeI This study fragment containing wild-type SNU71 was cut from pSNU71001 and cloned into the SmaI site of pRS315.
pSNU71015 Mutant snu71-101 in pRS315. Mutant allele This study was moved as the SphI/BssHII fragment from pSNU71003 into same sites of pSNU71011.
pSNU71016 Wild-type SNU71 in pRS316. The BamHI/SalI This study fragment from pSNU71011 was moved to the same sites of pRS316.
pSNU71017 Mutant snu71-101 in pRS314. Mutant allele was This study moved from pSNU71015 into pRS314 using the BamHI and SalI sites.
(continued)
103 Table 2.2 (continued)
Plasmid Description Source pSNU71018 Wild-type SNU71 and wild-type YHC1 in pRS316. This study The YHC1 ORF was removed from the BamHI site of pYHC1005 and placed into the BamHI site of pSNU71016. pYHC1003 YHC1-2 allele (L13F) in pRS315. A 1-kb PCR Chang lab product was amplified by YHCP1+P3 primers using SUP2 genomic DNA as the template and cloned into the BamHI site of pRS315. pYHC1004 YHC1-1 allele (L13S)in pRS315. A 1-kb PCR Chang lab product was amplified by YHCP1+P3 primers using SUP13 genomic DNA as the template and cloned into the BamHI site of pRS315. pYHC1005 Wild-type YHC1 in pRS315. A 1-kb PCR product This study containing the wild-type YHC1 gene was amplified by YHCP1+P3 primers using wild-type genomic DNA as the template and cloned into the BamHI site of pRS315. pYHC1023 YHC1-HA in pRS416. A 1-kb BamHI WT YHC1 Chang lab fragment with an HA tag at the 3’ end from pYHC1005 was cloned into pRS416. pYHC1027 L13A mutation using pYHC1005 as template Chang lab pYHC1028 L13C mutation using pYHC1005 as template. Chang lab pYHC1029 L13D mutation using pYHC1005 as template. Chang lab pYHC1030 L13E mutation using pYHC1005 as template. Chang lab pYHC1031 L13G mutation using pYHC1005 as template. Chang lab pYHC1032 L13H mutation using pYHC1005 as template. Chang lab
(continued)
104 Table 2.2 (continued)
Plasmid Description Source pYHC1033 L13I mutation using pYHC1005 as template. Chang lab pYHC1034 L13K mutation using pYHC1005 as template. Chang lab pYHC1035 L13M mutation using pYHC1005 as template. Chang lab pYHC1036 L13N mutation using pYHC1005 as template. Chang lab pYHC1037 L13P mutation using pYHC1005 as template. Chang lab pYHC1038 L13Q mutation using pYHC1005 as template. Chang lab pYHC1039 L13R mutation using pYHC1005 as template. Chang lab pYHC1040 L13T mutation using pYHC1005 as template. Chang lab pYHC1041 L13V mutation using pYHC1005 as template. Chang lab pYHC1042 L13W mutation using pYHC1005 as template. Chang lab pYHC1043 L13Y mutation using pYHC1005 as template. Chang lab
105 CHAPTER 3
RESULTS
3.1 Investigation of Alterations in the U1-C Protein and the U1 snRNA that
Allow Bypass ofPRP28
Prp28p, a putative DEAD-box RNA helicase, has been proposed to promote the switch of U1 for U6 at the 5’ SS (Staley and Guthrie, 1999). Using genetic strategies to illuminate Prp28p’s function in splicing, our laboratory found that mutations in YHC1, the gene encoding the U1-C protein, were able to bypass the requirement for the normally essential Prp28p. Specifically, mutations altering position L13 to F or S allowed bypass. Position L13 is strictly
conserved among species and lies within a C2H2-type zinc finger motif. As two alterations of the same amino acid were found to bypass Prp28p, site-directed mutagenesis was used to alter L13 to every other amino acid. Despite the absolute conservation of an L at position thirteen, six of the nineteen altered
106 amino acids behaved like the wild-type protein (C, I, M, N, Q, V) while all of the other variants allowed bypass of Prp28p (Chen et al., 2001). Mutations disrupting other conserved residues in the zinc finger were not able to allow bypass of Prp28p, indicating that position L13 may be involved in an important interaction that is distinct from that in which the zinc finger participates (T.-H.
Chang, unpublished). My work extends the analysis of the altered U1-C proteins to gain a better understanding of how they may allow bypass of Prp28p.
3.1.1 Determination of in vivo and in vitro Splicing Defects in the prp28∆
YHC1-1 Strain
The initial characterization of the prp28∆ YHC1-1 strain had revealed a cold-sensitive growth defect when PRP28 was not present (Chen et al., 2001).
As the genetic alterations that likely caused this phenotype affected splicing factors, pre-mRNA splicing was predicted to be blocked at low temperatures in this strain. To verify this prediction, the splicing of actin transcripts in these cells was analyzed using primer extension assays. For this in vivo analysis, total RNA was collected from wild-type, prp28∆ YHC1-1 and the cold-sensitive prp28-102 strains. RNA was prepared from cells that had been grown at the permissive
107 temperature of 30˚C as well as from cells that had been shifted to 16˚C, the non- permissive temperature at which splicing defects were predicted in both the prp28∆ YHC1-1 and prp28-102 strains. Primer extension was then used to detect the actin transcripts present in these cells. At the permissive temperature of 30˚C, only mature mRNA was detected in all strains tested. After being shifted to low temperature, however, a unspliced pre-mRNA accumulates in the prp28-
102 and prp28∆ YHC1-1 strains, indicating that splicing is hindered at low temperatures (figure 3.1, lanes 5 and 6).
To further investigate the splicing defect found in this bypass suppressor strain, splicing extract was prepared from wild-type and prp28∆ YHC1-1 cells and used for in vitro splicing reactions. As the bypass strain exhibited a cold- sensitive growth phenotype and accumulated unspliced messages at low temperatures in vivo, it was predicted that splicing in the in vitro assays would also be stalled at low temperatures. Splicing is inhibited in the prp28∆ YHC1-1 strain at 16˚C but proceeds normally at the permissive temperature of 25˚C
(figure 3.2, lanes 3 and 4). It was noted, however, that splicing at 25˚C in the prp28∆ YHC1-1 extract is less efficient than that in the wild-type extract (figure
3.2, compare lanes 1 and 3). A time-course study comparing the extent of
108 splicing in wild-type and prp28∆ YHC1-1 extracts at 25˚C revealed that the splicing proceeds at a slower rate in the bypass mutant extracts (figure 3.3). This finding was not unexpected, as the prp28∆ YHC1-1 strain exhibits a slight growth defect even at 25˚C (Chen et al., 2001). In addition, no intermediate splicing products are found in the prp28∆ YHC1-1 reactions at 16˚C. This indicates that splicing is blocked before the first trans-esterification reaction, a finding that is consistent with the roles of U1-C and Prp28p in early spliceosome formation.
3.1.2 Recombinant His6-Prp28p Relieves Splicing Defects in vitro
I observed a cold-sensitive block to splicing both in vivo and in vitro in the prp28∆ YHC1-1 strain, results that were consistent with the observed growth phenotype. This strain is no longer cold-sensitive when an allele of PRP28 is present. In an effort to recapitulate this in vitro and further develop our
biochemical tools for splicing analysis, recombinant His6-Prp28p was purified for
use in in vitro splicing reactions (figure 3.4 A). The recombinant His6-Prp28 protein was also submitted for antibody production that was monitored by
Western analysis (figure 3.4 B). The recombinant His6-Prp28 protein released the block to splicing at 16˚C in both prp28-102 and prp28∆ YHC1-1 extracts
109 (figure 3.5). Splicing in the prp28-102 extract was rescued to the level of splicing seen at permissive temperatures (figure 3.5, compare lanes 1 and 3). Although splicing was also rescued in the prp28∆ YHC1-1 extract, it did not completely recover to the level of activity seen at permissive temperature (figure 3.5, compare lanes 4 and 6). This observation may indicate a residual defect due to the YHC1-1 mutation (see Discussion).
3.1.3 Alterations in the U1 snRNA Bypass PRP28
It was speculated that the role of Prp28p is to unwind the U1 snRNA/5’ SS duplex to help release the U1 snRNP from the 5’ SS during spliceosome assembly (Staley and Guthrie, 1999). Therefore, it seemed possible that the alterations in the U1-C protein may somehow weaken the U1/5’ SS association, thus eliminating the need for Prp28p to disrupt the duplex. This idea is also consistent with the observation that the YHC1-1 mutation could no longer allow bypass of PRP28 at low temperatures. Under these conditions, the U1 snRNA/5’
SS duplex would be thermodynamically re-stabilized, preventing bypass. If this hypothesis is correct, one would predict that U1 snRNA mutations that decreased base pairing with the 5’ SS of the pre-mRNA, thus weakening the U1/5’ SS
110 interaction, may also be able to bypass PRP28. Concurrently, mutations that resulted in an increase of base pairing should not allow bypass. This hypothesis was tested by introducing several plasmid-borne mutant alleles of SNR19, the gene encoding the U1 snRNA, into a prp28∆ strain carrying PRP28 on a counter- selectable URA3 plasmid (Siliciano and Guthrie, 1988). As expected, the C4U and C8U mutations in the U1 snRNA that are predicted to disrupt the base pairing with intron positions G1 and G5, respectively, dominantly bypass PRP28 while the U5G and U5A mutations that are predicted to strengthen the U1/5’ SS interaction do not (figure 3.6). Interestingly, I also found that the U1-U5C mutation also allows bypass of PRP28. Position U5 in the U1 snRNA is normally located opposite position U4 in the intron of the pre-mRNA. Although this is not a typical Watson-Crick base pair, a U-U interaction has been documented in other situations (Baeyens et al., 1995; Butcher et al., 1997; Lietzke et al., 1996;
Schroeder et al., 1996). My result therefore suggests that there may be an interaction between the U’s at positions 4 and 5 in the intron and the U1 snRNA, respectively, that may contribute to stability of the U1/5’ SS association. My finding that duplex-weakening alterations in the U1 snRNA are able to bypass
111 PRP28 supports our model that the bypass alterations in the U1-C protein may function by weakening the interaction between the U1 snRNP and the 5’ SS
(figure 3.7).
3.1.4 Investigating the Growth Phenotypes of the U1-C L13 Variants
Alteration of position L13 within U1-C to any amino acid other than C, I, M,
N, Q or V allowed Prp28p to become dispensable (Chen et al., 2001). In an attempt to acquire more information about the putative interaction which requires the participation of position L13 of U1-C, a more in-depth analysis of the pool of
U1-C L13 variants was conducted. As the substitution of different amino acids at position 13 might have varying impacts on this interaction, the growth phenotypes of the L13 bypass mutants were analyzed, both as functional replacements for wild type U1-C as well as when allowing the cell to survive without Prp28p. First, I determined the growth patterns of strains in which the mutant proteins serve as the only source of U1-C in an otherwise wild-type cell.
As predicted, all of the mutants that could not bypass Prp28p behaved just like
112 wild-type U1-C at all temperatures (data not shown), while several of the bypass mutants resulted in only minor growth phenotypes at high (37˚C) or low (16˚C) temperatures (Figure 3.8).
The growth patterns of strains containing the U1-C mutants that bypass
Prp28p were also tested to determine to what extent each mutant could bypass.
In the absence of Prp28p, all bypass mutant strains grow well at 30˚C but are both cold-sensitive and temperature-sensitive (figure 3.9). This finding appears to be consistent with our model in which the U1-C bypass suppressors result in a more relaxed U1/5’ SS interaction. At low temperatures, the complex is more stable and again requires Prp28p for ensuing rearrangements while at higher temperatures the complex may become too unstable or not form efficiently (Chen et al., 2001).
3.1.5 Verification of Splicing Defects in Transcripts that are Highly
Sensitive to prp28∆ YHC1-1
Although I found that splicing is affected in the U1-C bypass mutant both in vivo and in vitro, these experiments were limiting as they only tested actin transcripts. The recent innovation of global splicing analysis using microarray
113 technology has provided us with a new tool to gain a more complete understanding of the genome wide effects of the absence of PRP28 as allowed by the YHC1-1 bypass allele (Clark et al., 2002; Spingola et al., 1999). This new technique was employed to ask whether the splicing of a specific subclass of transcripts was particularly affected by the U1-C L13F bypass mutant (T.-H.
Chang, unpublished). Although the complete details of the microarray results will be reported elsewhere, here I present the validation of some of these data by
RT-PCR. Total RNA was prepared from prp28∆ YHC1-1 cells carrying either a
PRP28 plasmid or an empty vector that had been incubated at either 30˚C or
16˚C. RT-PCR was subsequently employed to amplify several candidate transcripts. Among the messages that were found to be poorly spliced during the microarray analysis were RPL36B and QCR9 (T.-H. Chang, unpublished).
RPL36B encodes a component of the large ribosomal subunit that binds the 5.8
S rRNA and QCR9 encodes a subunit of the ubiquinol cytochrome C oxidoreductase complex (Lee et al., 1983; Phillips et al., 1990). RT-PCR revealed a defect in the splicing of both RPL36B and QCR9 transcripts (figure
3.10). In the case of RPL36B, I observed a decrease in the level of mRNA and an increase in the level of pre-mRNA in the absence of PRP28 at 16˚C (figure
114 3.10 B, lane 9). Similar results were seen with QCR9, although a smaller defect is visible even at the permissive temperature of 30˚C when PRP28 is not present
(figure 3.10 A, lanes 8 and 9).
3.2 Prp8p Collaborates with Prp28p During the U1/5’ SS Rearrangements
Our bypass suppressor strain provided us with the unique opportunity to further explore the target of Prp28p and it’s importance during the rearrangements at the 5’SS. By completely eliminating Prp28p, a normally essential component of the spliceosome activation pathway, we were able to ask what other factors normally cooperate with Prp28p to drive spliceosome activation. To this end, I screened for mutations that cause synthetic-lethality with YHC1-1 mutation in the absence of PRP28. Such mutations would likely occur in factors that are normally required to assist Prp28p during spliceosome assembly and would therefore become absolutely critical in its absence.
Mutations compromising the ability of these factors to perform during splicing would therefore re-impose the requirement for PRP28 in the presence of the
YHC1-1 mutation.
115 3.2.1 prp8-501 is Synthetically Lethal with prp28∆ YHC1-1
A strain containing the YHC1-1 bypass suppressor mutation and carrying
PRP28 on a counter-selectable plasmid was subjected to UV mutagenesis in order to isolate strains that could no longer survive without PRP28 (figure 3.11).
Approximately 100,000 colonies were screened and a single candidate was identified. Genetic analysis demonstrated that the mutation causing synthetic lethality was located in a single gene and was recessive (data not shown, see
Materials and Methods). To identify the gene that had been mutated, a wild-type genomic library was introduced into the synthetic lethal strain and colonies that could again lose PRP28 were isolated. After plasmid recovery and sequencing, a plasmid containing a single open reading frame (ORF) encoding the splicing factor Prp8p was identified, suggesting that a mutation in PRP8 was responsible for the observed synthetic lethality.
Prp8p is a well-studied, evolutionarily conserved component of the U5 snRNP (Hodges et al., 1995; Lossky et al., 1987). Prp8p is an extremely large
260 Kd protein and is known to interact with other splicing proteins as well as both the 5’ and 3’ splice sites of the pre-mRNA (Reyes et al., 1999; Reyes et al.,
1996; Teigelkamp et al., 1995a; Umen and Guthrie, 1995a; Umen and Guthrie,
116 1995b). Due to its large size and extensive interactions with many splicing factors, Prp8p is thought to serve as a large platform that coordinates the rearrangements that activate the catalytic core of the spliceosome (Collins and
Guthrie, 1999; Dix et al., 1998; Siatecka et al., 1999; Teigelkamp et al., 1995b).
My result is consistent with the previously identified genetic interactions between
PRP8 and PRP28. Mutations in PRP8 can suppress cold-sensitive mutations in
PRP28, supporting the idea that Prp8p somehow assists Prp28p during splicing
(Kuhn et al., 2002; Strauss and Guthrie, 1991).
Gap repair was used to identify the mutation in PRP8 that caused synthetic lethality with prp28∆ YHC1-1 (see Materials and Methods). I identified a novel allele of PRP8, prp8-501, that contains two mutations leading to the amino acid changes I1825K and L1835F. Interestingly, these alterations fall into the C-terminal portion of Prp8p that also contains many other alterations found to affect various aspects of splicing (figure 3.12). Specifically, prp8-501 lies within a region of PRP8 that contains many mutations that affect pyrimidine tract recognition as well as mutations that suppress splicing defects due to splice site mutations (Collins and Guthrie, 1999; Umen and Guthrie, 1995a; Umen and
Guthrie, 1996). In addition, the prp8-201 allele that was found to suppress a
117 cold-sensitive mutation in the U4 snRNA changes amino acid 1861 which is located only 26 and 36 amino acids downstream of the alterations caused by the prp8-501 allele (Kuhn et al., 1999). Finally, this region of the human Prp8p was also found to cross link to the 5’ SS of the pre-mRNA, part of the proposed target of Prp28p (Reyes et al., 1999).
3.2.2 Mutations Throughout PRP8 are Synthetically Lethal with prp28∆
YHC1-1
Several previous studies of Prp8p revealed regions of the protein that appeared to be required for specific tasks during splicing. I was therefore interested in knowing whether a particular region of Prp8p may be required to assist Prp28p during spliceosome activation. To address this question, I took advantage of the large pool of PRP8 alleles previously identified in other labs and asked whether any of these, particularly those located near prp8-501, could also re-impose the requirement for PRP28 in the presence of the YHC1-1 bypass mutation. The specific mutations investigated and the result of each test are listed in table 3.1. Alleles throughout the entire PRP8 ORF were tested, but only
118 those causing alterations within a small region of approximately 300 amino acids within Prp8p (1574 to 1861) were synthetically lethal with prp28∆ YHC1-1 (figure
3.13).
3.3 Identification and Preliminary Characterization of prp42-101 and snu71-
101 as Bypass Suppressors of PRP28
To gain a better understanding the function of Prp28p, our lab has focused on defining precisely what constitutes the in vivo substrate of Prp28p. As discussed above, we were able to show that the U1-C protein is likely to be included in the target of Prp28p, indicating that this protein may function as an
RNPase (Chen et al., 2001). This exciting possibility prompted us to ask whether additional proteins might be included in Prp28p’s substrate.
Formation of the U1/5’ SS duplex is critical as it is responsible for initiating splicing of a transcript. It therefore seems unlikely that the short 6-base pair U1 snRNA/5’ SS duplex and the U1-C/5’ SS association are the only interactions that mediate such an important step. It was also found that U1 snRNPs are still able to bind to transcripts, albeit with low efficiency, in extracts depleted of U1-C, indicating that additional factors may be involved in stabilizing the interaction
119 (Heinrichs et al., 1990). As the yeast U1 snRNP contains at least 17 proteins, it seemed very likely that some of these factors may also play a role in stabilizing this interaction, particularly those that have been found to cross-link to the 5’ SS region (Zhang and Rosbash, 1999). We also appreciated the fact that novel, potentially non-U1 proteins that may not be easily identified by biochemical purification or cross-linking experiments due to weak or transient interactions may still remain unidentified. We predicted that like U1-C, any additional factors that may normally help to stabilize the U1/5’ SS interaction may also be counteracted by Prp28p. In theory, specific alterations in these proteins should also weaken the U1/5’ SS interaction and allow bypass of Prp28p.
To detect additional protein targets of Prp28p, other members of our lab conducted a genetic screen to identify non-YHC1 bypass suppressors of Prp28p.
The screen yielded recessive mutations in three additional genes. Three different mutations were identified within YNL187W, a previously uncharacterized non-essential gene (R. Hage, R. Pengal, S. Hamzehzadeh, and T.-H. Chang, unpublished). Preliminary studies revealed that these mutations are likely loss- of-function as the deletion of YNL187W also allows bypass. Interestingly,
Ynl187Wp was found to interact with the SmB protein in two-hybrid screens,
120 indicating it may be a splicing factor. Studies on this gene are continuing to explore its possible role in splicing (R. Hage and T.-H. Chang, unpublished).
Here I present the identification of the two additional genes and the mutations within them that allow bypass of PRP28. One strain obtained from the screen contains a novel allele of PRP42 which we have designated prp42-101.
While its specific function in splicing has not be studied in great detail, Prp42p is an essential splicing factor known to be a yeast-specific U1 snRNP component
(Gottschalk et al., 1998; McLean and Rymond, 1998; Rigaut et al., 1999).
Although Prp42p was not initially identified in purified yeast U1 snRNP particles, later studies found that it is a weakly associated component and that a mutation in PRP42 is synthetically lethal with mutations in the U1 snRNA (Gottschalk et al., 1998; Neubauer et al., 1997; Rigaut et al., 1999). In addition, the U1 snRNP adopts an altered conformation and is defective in binding to the pre-mRNA in extracts prepared from cells metabolically depleted of Prp42p (McLean and
Rymond, 1998).
The investigation of Prp42p as a putative splicing factor was initiated upon its identification in a search for proteins with sequence similarity to Prp39p, another yeast U1 snRNP protein. The amino acid sequence of Prp42p is 50%
121 similar to that of Prp39p, much of which is due to analogous clusters of tetratricopeptide repeat (TPR) motifs (McLean and Rymond, 1998). A large number of proteins containing TPR repeats have been found in organisms ranging from bacteria to humans and are implicated in many cellular processes including cell cycle control, stress response and transcription repression. The
TPR motif has been defined as a 34-amino acid repeat, although some variations and degeneracies are found in certain sub-families of these proteins. In general,
TPR motifs are thought to form amphipathic α-helices that are likely to be involved in protein-protein interactions. The motif is divided into domains A and
B that physically interact with the opposing domains of other TPR repeats to create a “snap helix”. Domain A contains a conserved consensus sequence (W-
LG-Y) that forms a hydrophobic pocket while domain B contains a bulky phenylalanine side chain (A-F-A) that fits into this pocket, forming a “knob-in- hole” interaction (Lamb et al., 1995). Some TPR proteins contain stretches of consecutive repeats, and these knob-in-hole interactions are thought to allow stacking of the helices to form complex intramolecular structures (Goebl and
Yanagida, 1991). In other cases, these motifs are involved in interactions with another TPR protein or with helical domains of non-TPR proteins (Lamb et al.,
122 1995). It has been suggested that the TPR motifs of Prp39p and Prp42p may allow these to factors to interact with one another or may be important for other protein-protein contacts within the U1 snRNP (Gottschalk et al., 1998).
The consensus sequence of the TRP motifs in Prp39p and Prp42p are most closely related to those found in the Drosophila crooked neck (Crn) protein, a factor required during embryogenesis (Zhang et al., 1991). Specifically, the domain A structures of Prp39p and Prp42p are very consistent with those of Crn while the domain B structures are less conserved but still retain a general TPR consensus and an overall alpha-helical character. A database search for proteins containing TPRs with domain A sequences of the Prp39p-Pp42p-Crn type revealed a number of proteins that are part of the RNA cleavage stimulation factor required for pre-mRNA 3’ end processing. These factors include the yeast
Rna14p, Drosophila suppressor of forked [Su(f)] and human CstF77. When combined with Prp39p and Prp42p, these factors may constitute a putative sub- family of TPR proteins involved in RNA metabolism (McLean and Rymond,
1998). Although the specific function of the Drosophila Crn protein is unclear, the yeast homologue of this protein has been identified as Clf1p and appears to
123 assist in stabilizing the U4/U6/U5 tri-snRNP interaction with the spliceosome, indicating an RNA-processing role for this factor as well (Chung et al., 1999).
The second strain from the bypass suppressor screen was found to contain a novel allele of SNU71 which we have designated snu71-101. Like
Prp42p, little is know about Snu71p other than it is an essential component of the yeast U1 snRNP that does not have an obvious counterpart in the mammalian
U1 snRNP (Gottschalk et al., 1998). Interestingly, Snu71p contains 19 RS-, RD- or RE-dipeptides similar to those found in the metazoan family of SR-proteins. In contrast to the metazoan SR-dipeptides which are usually contained within a single protein domain, the SR-dipeptides within Snu71p are spread throughout the protein (Gottschalk et al., 1998). As discussed in the introduction, mammalian SR proteins appear to play a role in splice site selection and can bypass the requirement for the U1 snRNP when over-expressed (Crispino et al.,
1994; Tarn and Steitz, 1994). Snu71p is an integral component of the U1 snRNP. Antibodies against Snu71p co-precipitate the U1 snRNA in the presence of 700 mM salt, indicating that Snu71p has a much stronger association with the
U1 snRNP than does Prp42p, which only co-precipitates the U1 snRNA in100 mM salt (Gottschalk et al., 1998; McLean and Rymond, 1998). Interestingly,
124 Snu71p may also be involved in interactions with the cap binding complex (CBC) as it was isolated in a genetic screen seeking mutants that caused lethality in the absence of the CBC (Fortes et al., 1999). A connection with the CBC may also support a role for Snu71p at the 5’ SS as interactions between the CBC and U1 snRNP factors are known to affect 5’ SS choice (Lewis et al., 1996).
3.3.1 Novel Alleles of PRP42 and SNU71 Bypass the Requirement for
Prp28p
Before the bypass genes were identified in the remaining two bypass suppressor candidate strains, they were first back-crossed to decrease the number of plasmids present to facilitate analysis. The resulting strains were then transformed with a wild-type yeast genomic library. As the bypass suppressor mutations were recessive, the presence of a wild-type copy of the mutated gene would reinstate the requirement for PRP28. Library transformants that lost their bypass activity, i.e. they again became dependent on the presence of a wild-type
PRP28 plasmid, were identified. Following plasmid rescue and verification that the resulting plasmid was responsible for loss of bypass activity, plasmids were subjected to sequencing and a BLAST search to identify the open reading frames 125 (ORFs) present. The plasmid recovered from one candidate contained several
ORFs including PRP42. As this was the most likely to be our gene of interest, the PRP42 ORF was sub-cloned from the library plasmid into yeast expressions vectors to verify it’s identity as the bypass gene. As predicted, the presence of wild-type PRP42 was sufficient to eliminate the bypass ability of the strain (data not shown). The library plasmid from the second candidate strain also contained several ORFs, and again the obvious candidate SNU71 was identified. After sub-cloning the SNU71 ORF into yeast expression vectors, a plasmid containing only SNU71 was sufficient to eliminate the bypass activity of the second candidate, confirming that a mutation in SNU71 could bypass PRP28 (data not shown).
The mutations within PRP42 and SNU71 responsible for bypassing
PRP28 were identified by sequence analysis of the mutant alleles amplified by the polymerase chain reaction (PCR) using genomic DNA from the bypass strains as templates. A single guanine to thymine change predicting a D76Y amino acid alteration was identified within PRP42, and we have designated this allele prp42-101. The alteration lies within the first TPR protein-protein interaction domain (figure 3.14). Analysis of the mutant SNU71 locus revealed a
126 single base change at T 1410 to C, predicting a change of leucine to serine change at amino acid 270. This alteration lies between regions encoding two of the SR-like dipeptides within Snu71p. We have designated this allele snu71-101
(figure 3.15).
3.3.2 The prp28∆ prp42-101 Strain Displays a Cold-sensitive Growth
Phenotype
During the identification and isolation of candidates from secondary the bypass suppressor screen it was noted that the new strains appeared to be cold- sensitive in the absence of PRP28, much like the original prp28∆ YHC1-1 bypass strain (R. Hage and T.-H. Chang, unpublished). As these strains had been extensively mutagenized by UV light during the screen, it was not initially clear whether the cold-sensitive growth phenotypes were linked to the bypass mutation or whether additional mutations were the source of the growth defects. To resolve this issue, I back-crossed the candidate strains later found to contain the prp42-101 and snu71-101 alleles to the non-mutagenized parent strain three times in order to eliminate extraneous mutations (see Materials and Methods).
After each mating the phenotypes of the progeny were followed and it appeared
127 that the cold-sensitive phenotype did co-segregate with the bypass alleles (data not shown). After the final back-cross the growth phenotype of the prp28∆ prp42-101 strain was more precisely analyzed using a spot test to compare the growth of this strain with and without PRP28 to the growth of the parental strain used for the screen. In the presence of a plasmid carrying wild-type PRP28 the prp28∆ prp42-101 strain grows much like the parental strain at all temperatures.
In the absence of PRP28, however, the prp28∆ prp42-101 strain shows a slightly slower growth rate at 30˚C, a more severe defect at 25˚C, and an inability to grow at 16˚C (figure 3.16 A).
3.3.3 prp42-101 Does not Cause a Growth Defect on its Own
Although there appeared to be no growth phenotype in the prp28∆ prp42-
101 strain when a PRP28 plasmid was present, the prp42-101 mutation was isolated in a strain that had never been subjected to mutagenesis and carried a wild-type copy of PRP28 on the chromosome to ensure that there was no growth phenotype associated with this allele. Plasmids bearing either PRP42 or prp42-
101 were introduced into a strain in which the chromosomal copy of PRP42 was deleted and all other loci were wild-type. A spot test was then used to compare
128 the growth phenotypes of these strains at various temperatures. Consistent with earlier findings, this test revealed that although there may be a very slight growth defect at 37˚C there are no discernable differences in cell growth at any other temperature (figure 3.16 B).
3.3.4 Deletion of prp42 Cannot Bypass PRP28
For several of the bypass suppressors that have been identified to date, it has been found that the deletion of one gene can bypass the requirement for another. In the case of the splicing factors MUD2 and SUB2, the deletion of
MUD2 can bypass the requirement for SUB2 (Kistler and Guthrie, 2001). In addition, the deletion of YNL187W can bypass PRP28 (R. Hage and T.-H.
Chang, unpublished). These findings prompted us to ask whether the deletion of
PRP42 can bypass PRP28. We predicted that this was unlikely to be the case as PRP42 is an essential gene while MUD2 and YNL187W are both non- essential. Nevertheless, prp42-101 is a recessive, presumably loss of function mutation and I therefore tested whether the complete deletion of PRP42 could also bypass PRP28. A strain with chromosomal deletions of both PRP28 and
PRP42 that carried a counter-selectable URA3 plasmid with wild-type copies of
129 both genes was used for this test (see Materials and Methods). Cells cannot survive in the absence of the plasmid carrying the wild-type genes, indicating that deletion of PRP42 cannot bypass the requirement for PRP28 (figure 3.16 C).
3.3.5 Analysis of the in vitro Splicing Defect in prp28∆ prp42-101
As the prp28∆ prp42-101 strain was cold-sensitive, we predicted that splicing of artificial actin transcripts in vitro would be blocked before the first catalytic step of splicing at low temperatures. Splicing extract was prepared from the prp28∆ prp42-101 bypass strain and tested for the ability to splice actin transcripts under at 25˚C and 16˚C. Surprisingly, I found that although splicing was inhibited at low temperatures, I did not see the complete block to splicing that we had expected (data not shown). To better characterize the defect, a time course study comparing splicing in the prp28∆ prp42-101 extract to that in a wild- type extract was conducted. This study revealed that while splicing is not completely prevented in the bypass extract at 16˚C, it is slowed (figure 3.17).
130 3.3.6 Synthetic Lethality of prp42-101 and snu71-101 with YHC1 Bypass
Alleles
The cold-sensitive phenotypes of the prp42-101 and snu71-101 bypass strains led us to propose that like U1-C, Prp42p and Snu71p may normally help to stabilize the U1/5’ SS interaction, and that alteration of these proteins may lead to a weakened association. This model predicts a unified function for all of these bypass factors (i.e. U1-C, Prp42p, Snu71p and Ynl187wp) at the stage of splicing requiring Prp28p. It was therefore predicted that combining two of the bypass alleles in one strain may cause an insurmountable defect that could prove lethal to the cell. More specifically, combining two mutations that both destabilize the U1/5’ SS interaction may lead to an extremely loose, non- functional duplex or could possibly prevent the association from occurring. To test this idea, the prp42-101 and snu71-101 bypass alleles were combined with the alleles of YHC1 that also bypass PRP28 to determine whether this would cause synthetic-lethality. Diagrams showing the approach used to test whether the prp42-101 and snu71-101 mutations were synthetically lethal with the YHC1 bypass alleles as well as the results of these tests are shown in figures 3.18 and
3.19 respectively. As expected, the bypass alleles of YHC1 were indeed
131 synthetically lethal with both prp42-101 and snu71-101 while the non-bypass alleles were not. Only minor exceptions were found; the mutation causing the
U1C L13M alteration that does not allow bypass was synthetically lethal with prp42-101, and the mutation causing the U1C L13Q alteration showed weaker growth than expected when combined with snu71-101.
3.3.7 Alleles of prp8 are Synthetically Lethal with prp28∆ prp42-101
In section 3.2.2 mutations mapping to a small region of the highly conserved splicing factor PRP8 were shown to be synthetically lethal with prp28∆
YHC1-1. As my synthetic-lethality data indicate that the prp42-101 mutation may be functioning in a similar manner as the YHC1-1 mutation, I tested whether the prp42-101 mutation also demonstrated genetic interactions with PRP8. A strain was constructed that lacked PRP28, carried the prp42-101 bypass allele and allowed the introduction of the numerous PRP8 alleles by plasmid shuffling (see
Materials and Methods). The results of this test are shown in figure 3.20 and are listed in greater detail in table 3.1. Although there were several differences, I found that the mutations that were synthetically lethal with prp42-101 map to the same region of PRP8 as those that are synthetically lethal with YHC1-1.
132 3.4 Bypass of PRP28 is a Specific Event
Preliminary findings from secondary bypass suppressor screen as well as data from two additional studies prompted us to directly investigate several U1 snRNP proteins for the ability to bypass Prp28p. First I became interested in asking whether specific alterations in the SmB, SmD1 and SmD3 proteins may allow bypass of Prp28p. The U1 snRNP contains the common set of seven Sm proteins that form a heteromeric ring that is also found in the U2, U4 and U5 snRNPs (Yu, 1999). Three of these proteins, SmB, SmD1 and SmD3, are adjacent to one another in the order D1-B-D3 within the Sm ring and cross-link to the pre-mRNA close to the 5’ SS (Hermann et al., 1995; Kambach et al., 1999;
Lehmeier et al., 1994; Raker et al., 1999; Zhang and Rosbash, 1999).
Specifically, SmB, SmD1 and SmD3 stabilize the interaction between the U1 snRNP and the pre-mRNA through their highly charged C-terminal tails. Zhang et al. (2001) found that when the C-terminal tails of these Sm proteins are truncated, formation of the U1 snRNP/pre-mRNA complex is impaired. Yeast strains containing tail truncated versions of two of these proteins cause either temperature-sensitivity or lethality, consistent with their defect in commitment complex formation. This study shows that that like U1-C, the SmB, SmD1 and
133 SmD3 proteins are in contact with the pre-mRNA and are required for the U1 snRNP to form a stable association with the 5’ SS. Although this result alone indicated that these three Sm proteins could potentially part of the target of
Prp28p, a second study supported this idea by showing that they are also located very near the U1-C protein. The three-dimensional structure of the human U1 snRNP as determined by electron cryomicroscopy revealed that the U1-C protein is located directly above these three Sm proteins, consistent with a previously identified cross link between human U1-C and SmB proteins in purified HeLa U1 snRNPs (Nelissen et al., 1994; Stark et al., 2001). Taken together, these findings introduced the possibility that U1-C may have an indirect influence on the U1/5’ SS interaction. Specifically, the U1-C protein may make critical contacts with the Sm proteins that help to position them properly, thus allowing their charged tails to interact with the pre-mRNA. The alteration of position L13 within U1-C could alter this putative interaction, leading to reduced interplay between the charged Sm tails and the pre-mRNA. The subsequent weakening of the U1/5’ SS duplex could then in turn bypass Prp28p. If this were indeed the case, directly altering the Sm proteins in a way that disrupts their interaction with the pre-mRNA would be predicted to bypass Prp28p. I therefore tested whether
134 Prp28p could be bypassed by the tail truncated versions of the Sm proteins B,
D1 and D3 that hinder commitment complex formation.
In addition to testing the Sm proteins, the first candidate gene identified in the secondary bypass suppressor screen prompted us to directly test two more
U1 snRNP proteins for bypass activity. Mutations in YNL187W, a non-essential gene, can bypass PRP28. Furthermore, deletion of this gene also allows bypass
(R. Hage and T.-H. Chang, unpublished). Although Ynl187wp has not been shown to associate with the U1 snRNP, it may be a loosely associated factor that was missed during the previous biochemical fractionations. This possibility is supported by our findings that all of the other bypass suppressors of Prp28p including U1-C, Prp42p and Snu71p are all known U1 snRNP components
(Gottschalk et al., 1998; McLean and Rymond, 1998; Neubauer et al., 1997;
Rigaut et al., 1999). We noted that like YNL187W, two genes encoding known
U1 snRNP proteins, MUD1 and NAM8, are also non-essential (Ekwall et al.,
1992; Liao et al., 1993). The involvement of NAM8 was investigated by another member of the lab and will be reported in full elsewhere (R. Hage and T.-H.
Chang, unpublished). MUD1 encodes Mud1p, the yeast homologue of the mammalian U1-A protein and was initially identified in a screen for factors that
135 were synthetically lethal with mutations in the U1 snRNA. Mud1p contains two conserved RNA binding domains (RBD), one of which binds to the U1 snRNA.
While the target of the other RBD is unclear, it is required for splicing (Liao et al.,
1993; Tang and Rosbash, 1996). Mud1p is able to co-precipitate the U1 snRNA and biochemical purifications confirmed its presence in the U1 particle
(Gottschalk et al., 1998; Liao et al., 1993; Neubauer et al., 1997; Rigaut et al.,
1999). Although it seemed unlikely, we decided to test whether the deletion of the genes encoding these non-essential U1 proteins could bypass PRP28. As these proteins are integral components of the U1 particle, their deletion could disrupt the U1 snRNP and result in a weakened association with the 5’ SS, thus allowing bypass of PRP28.
3.4.1 Truncation of SmB, SmD1 and SmD3 C-terminal Tails Cannot Bypass
Prp28p
Strains that combined the tail truncated versions of the Sm proteins with
PRP28 on a counter-selectable plasmid were tested for the ability to survive in the absence of the PRP28 plasmid. The tail truncated versions of SmB, SmD1 or SmD3 were not able to survive in the absence of the PRP28 plasmid,
136 indicating that these alterations could not bypass Prp28p. Similarly, even when the tail truncated versions of both SmB and SmD3 were present, a combination that causes a temperature-sensitive growth phenotype, the PRP28 plasmid was still required for cell viability (figure 3.21).
3.4.2 Deletion of MUD1 Cannot Bypass PRP28
To test whether deleting MUD1 could bypass Prp28p, a MUD1 deletion strain was purchased and crossed to our strain that carried PRP28 on a counter- selectable plasmid. Following tetrad dissection I found that progeny lacking
MUD1 could not survive without the PRP28 plasmid. Figure 3.22 shows four such progeny streaked to selective plates to confirm their inability to bypass
PRP28. The deletion of NAM8 was also unable to bypass PRP28 (R. Hage, personal communication).
137 30˚C 16˚C
1 2 3 4 5 6
1,353 1,078 872
603
310
Figure 3.1. Primer extension analysis of splicing in prp28∆ YHC1-1 cells reveals a splicing defect at 16˚C. Splicing of actin transcripts was analyzed in PRP28 (lanes 1 and 4), prp28-102 cold-sensitive (lanes 2 and 5) and prp28∆ YHC1-1 (lanes 3 and 6) strains by primer extension. Total RNA was collected from cells grown at 30˚ C or grown at 30˚C and then shifted to 16˚C for 16 hours. The positions of the cDNA products is indicated on the right, and positions of a DNA marker are indicated in nucleotides on the left.
138 Figure 3.2. In vitro splicing in prp28∆ YHC1-1 extracts. Splicing reactions were assembled using either wild-type or prp28∆ YHC1-1 extracts and incubated at 16˚C for one hour or 25˚C for 20 minutes. Diagrams illustrating the splicing products are shown to the right.
139 prp28∆ WT YHC1-1
25˚ 16˚ 25˚ 16˚
140 Figure 3.3. Time course study of splicing at 25˚C in wild-type and prp28∆ YHC1- 1 extracts. Splicing reactions were incubated at 25˚C for the number of minutes noted at the top of each lane. Diagrams showing the splicing products are shown to the right.
141 Wild-type prp28∆ YHC1-1
0 3 7 15 30 60 0 3 7 15 30 60
142 A 1 2 3 4 5 6B 1 2 3 190
120
97 85
68 60
50 43
29 40
18
Figure 3.4. Purification of recombinant His6-Prp28p and analysis of antibody production. A) SDS-PAGE analysis of pre-induction (lane 1), IPTG induction (lane 2), supernatant from PEI precipitation (lane 3), ammonium sulfate precipitate (lane 4), heparin column eluate (lane 5) and nickel column eluate (lane 6). Approximately 5 µg of protein from each fraction was separated on an 8% polyacrylamide (29:1) gel that was then silver stained. The positions of molecular weight standards in kilodaltons (Kd) are indicated on the left. B) Anti-
His6-Prp28p antibody. Western analysis of 100 µg of yeast total protein from strains with wild-type PRP28 (lane 1) and overexpressing PRP28 from a GPD promoter (lane 3). Recombinant His6-Prp28p is shown in lane 2. The anti- Prp28p serum was used at a 1:20,000 dilution for Western analysis.
143 prp28∆ prp28-102 YHC1-1
1 2 3 4 5 6
Figure 3.5. Recombinant His6-Prp28p rescues splicing defects in vitro. Lanes 1- 3 are reactions in prp28-102 cold-sensitive extracts while reactions in lanes 4-6 are in prp28∆ YHC1-1 extracts. Reactions were incubated at 25˚C (lanes 1 and 4) or 16˚C in the absence (lanes 2 and 5) or presence (lanes 3 and 6) of 6 pmol recombinant His6-Prp28p protein. Diagrams of the splicing products are shown on the right.
144 Figure 3.6. Mutations in the U1 snRNA can bypass PRP28. A) Base pairing between the U1 snRNA and the pre-mRNA 5’ SS. Bases in the U1 snRNA are numbered from its 5’ end. Exons in the pre-mRNA are boxed. B) Specific mutations in the U1 snRNA bypass PRP28. Strains containing various mutant U1 clones were spotted to 5-FOA plates and incubated at 30˚C. Serial dilutions of cell cultures are shown from the top down. Adapted from Chen et al., 2001.
145 A U1
Prp28p UCCAUUCAUA
UGGUAUGUUC UGGUAUGUUC
UAGAGACAUA U6
B U1
U1-C*
UCCAUUCAUA
UGGUAUGUUC UGGUAUGUUC
UAGAGACAUA U6
Figure 3.7. Model for bypass of Prp28p by alterations in U1-C. A) In the wild- type situation, the U1 snRNA/5’ SS interaction is disrupted by Prp28p, allowing the U6 snRNA/5’ SS interaction to occur. B) In the presence of altered U1-C proteins, the U1/5’ SS association is weakened, allowing this interaction to be replaced by the U6/5’ SS interaction in the absence of Prp28p.
146 Figure 3.8. Growth phenotypes of the YHC1 L-13 bypass mutants in a yhc1∆ strain. Bypass mutant alleles of YHC1 were introduced into a yhc1∆ strain by plasmid shuffling to determine their ability to serve as the only source of U1-C in an otherwise wild-type cell. The temperature of incubation is listed above each panel.
147 16°C 25°C 30°C 37°C
WT A D E F G H K P R S T W Y
148 Figure 3.9. Growth phenotypes of the YHC1 L-13 bypass mutants during bypass. The mutant alleles of YHC1 were introduced into a prp28∆ strain carrying PRP28/URA3 plasmid and spotted to 5-FOA to determine how efficiently they allow bypass of Prp28p. The temperature of incubation is listed above each panel.
149 16˚C 25˚C 30˚C 37˚C
A D E F G H K P R S T W Y
150 A 1 2 3 4 5 6 7 8 9 10 1,000 850 650 500 400 300 200
100
B 1 2 3 4 5 6 7 8 9 10 1,000 850 650 500 400 300 200
100
Figure 3.10. Agarose gel analysis of RT-PCR products to confirm splicing defects identified during microarray analysis. A) RT-PCR of QCR9 messages. B) RT-PCR of RPL36B messages. For both panels A and B: the first four lanes are control reactions of the un-spliced U1 snRNA and lanes 5 and 10 contain a DNA ladder, the sizes of the bands are shown to the right in base pairs. The RNA used for RT-PCR was collected from PRP28 YHC1-1 (lanes 1, 2, 6 and 7) or prp28∆ YHC1-1 (lanes 3, 4, 8 and 9) cells that were grown at 30˚C (lanes 1, 3, 6 and 8) or grown at 30˚C and then shifted to 16˚C (lanes 2, 4, 7 and 9).
151 Figure 3.11. Diagram of screen to identify mutations synthetically lethal with prp28∆ YHC1-1. A) Cells containing a PRP28 plasmid that can be lost due to the YHC1-1 mutation were B) exposed to UV light to induce mutations. C) In the presence of the mutation causing synthetic lethality the PRP28 plasmid must be retained. D) These cells were transformed with a wild-type genomic library, allowing the identification of cells that can again lose the PRP28 plasmid. E) Plasmid rescue and sequencing allow the identification of the gene PRP8. F). Gap repair was used to recover the mutation causing synthetic lethality, prp8- 501.
152 A B C prp28 prp28 prp28 ∆ UV ∆ ∆ YHC1-1 YHC1-1 YHC1-1
PRP28 PRP28 PRP28 x
PRP28 plasmid can be lost Cells are exposed to UV light PRP28 plasmid must be maintained in due to YHC1-1 mutation to induce random mutations presence of synthetic lethal mutation 153
F E D prp28 ∆ prp8-501 X PRP8 YHC1-1
= X x
Gap repair allows the identification of Library plasmid is recovered and Library plasmid with wild-type gene X the novel mutation within PRP8 sequenced to reveal that gene X is PRP8 allows PRP28 plasmid to again be lost Figure 3.12. Location of altered amino acids due to prp8-501 mutations. The numbers at the top of the diagram represent amino acid positions. The dotted yellow lines indicate the position of the alterations resulting from the prp8-501 allele. The small red box represents a domain identified as important for 3’ splice site fidelity. The large blue box represents the region containing mutations that can suppress splice site mutations. The black line at position 1861 represents the mutation found to suppress the U4-cs1 allele; white boxes represent the positions of the suppressors of this allele identified in a secondary screen. The lightning bolt indicates the position of the cross-link between the hPrp8 protein and the 5’ splice site. Black brackets indicate putative intra-molecular folds within the protein.
154 1964-1968
1 236 362 611 684 788 861 1094 1197 1371 1608 1817 1825 1835 1861 2065 2413 155
Prp8p Figure 3.13. Mutations altering a distinct region of Prp8p are synthetically lethal with prp28∆ YHC1-1. Features of the Prp8p map are as in figure 1.9. Mutations throughout PRP8 were tested for synthetic lethality with prp28∆ YHC1-1. Alterations that caused synthetic lethality map to an approximately 300 amino acid region of Prp8p. Dotted lines represent the amino acid changes caused by prp8-501 and solid lines represent amino acid alterations that also caused synthetic lethality.
156 1964-1968
1 236 362 611 684 788 861 1094 1197 1371 1608 1817 1861 2065 2413 157
D1700G E1832A,V W1608R L1794F N1883D
W1574R Q1677F, D1778G T1861P D1700E I1825K, L1835F Table 3.1. PRP8 mutations that are synthetically lethal with prp28∆ YHC1-1and prp28∆ prp42-101. The alteration(s)/alleles tested, their phenotype(s) determined in previous studies and the result of synthetic lethality tests with prp28∆ YHC1-1 and prp28∆ prp42-101 are listed. Abbreviations are: cs = cold sensitive, ts = temperature sensitive, ND = not determined, SL = synthetically lethal, y = yeast, h = human, Pyr defect = defective in pyrimidine tract recognition, SS Sup = splice site suppressor (both 5’ and 3’), Sup U4-cs1 = suppressor of U4-cs1, Sup U4-cs1, 2° = suppressor of U4-cs1 identified in secondary screen, Hyper U1 = exacerbates hyperstabilized U1/5’SS defect, 5’SS Sup = Suppressor of 5’ splice site mutations, Not 5’SS Sup = not a suppressor of 5’ splice site mutations
158
Synthetically Lethal? Alteration(s)/ allele Phenotype YHC1-1 prp42-101 I1825K, L1835F(prp8-501) SL prp28∆ YHC1-1 YES YES E1960K (prp8-101) cs, Pyr defect, SL prp16-301 SICK YES E1960G (prp8-102) Pyr defect no SICK F1834L (prp8-103) Pyr defect no no F1834S (prp8-104) Pyr defect no no W1608R (prp8-121) SS Sup, Hyper U1 YES YES W1574R (prp8-122) SS Sup YES no E1575V (prp8-123) SS Sup no no T1564A (prp8-125) SS Sup nono M1397I (prp8-124) SS Sup, Hyper U1 no YES E1817G (D-135) SS Sup noSICK N1869D (D-136) SS Sup no no K1864E (D-143) SS Sup no no F2176S, Q2313R, T2346A(prp8-144) SS Sup no no R236G Sup U4-cs1, 2° no no L280P Sup U4-cs1, 2°, sup prp28-1 no no K611R Sup U4-cs1, 2° no no E788G Sup U4-cs1, 2° no no D1094A Sup U4-cs1, 2° no no T1861P (prp8-201) ts, Sup U4-cs1, Hyper U1 YES YES N1809D Sup U4-cs1, 2° no no I1851L Sup U4-cs1, 2° no no T1702I not 5’SS Sup no no M1783K not 5’SS Sup no no I1933F not 5’SS Sup no no L1794F not 5’SS Sup YES YES S1906G not 5’SS Sup no no D1670N, D1778G not 5’SS Sup no ND (continued) Table 3.1 PRP8 Mutations Synthetically Lethal with prp28∆ YHC1-1and prp28∆ prp42-101
159 Table 3.1 (continued)
Synthetically Lethal? Alteration(s)/ allele Phenotype YHC1-1 prp42-101 D1700G not 5’SS Sup YES YES N1883D not 5’SS Sup YES YES S1769S, D1778G not 5’SS Sup no YES I1775T, D1778E not 5’SS Sup YES YES D1778G, N1869S not 5’SS Sup no SICK S1745P, D1778G not 5’SS Sup YES YES D1778R, L1779M, G1635D not 5’SS Sup YES YES S1829C, E1832V not 5’SS Sup YES YES E1832A, L1891P not 5’SS Sup YES YES E1832V, P1952L not 5’SS Sup no YES K1807R, S1829C, E1832A not 5’SS Sup YES YES Q1677R, D1700E not 5’SS Sup YES YES S1745F not 5’SS Sup no no F1724L not 5’SS Sup no no E1842N not 5’SS Sup no no N1792S, E1842N not 5’SS Sup no no Y1787C, E1842N not 5’SS Sup no no N1869D 5’SS Sup no no N1869D, S1970R (prp8-152) 5’SS Sup no no S1970R 5’SS Sup no no T1982A (prp8-153) 5’SS Sup no no T1982A, SA1966/7 AG (prp8-154) 5’SS Sup no no T1982A, V1987A (prp8-155) 5’SS Sup no no N1869D, T1982A (prp8-156) 5’SS Sup no no N1869D,S1970R, T1982A,V1987 not 5’SS Sup no no y1962-1975 to h1890-1903 not 5’SS Sup no no
160 Figure 3.14. An alteration in Prp42p allows bypass of Prp28p. Blue boxes indicate putative TPR motifs, the amino acid positions are noted at the top of the map. The dotted line indicates the position of the D76Y alteration caused by the prp42-101 mutation found to bypass Prp28p.
161 1 71 104 106 139 144 177 276 309 310 339 544 162
Prp42p Figure 3.15. An alteration in Snu71p allows bypass of Prp28p. The protein contains eight SR repeats shown as grey lines and nine SR-like repeats shown as blue lines. The black dotted line indicates the position of the L270S alteration caused by the snu71-101 mutation that can bypass Prp28p.
163 1 620 164
Snu71p Figure 3.16. Growth phenotype of strains containing prp42-101. A) Growth of the prp28∆ prp42-101 strain in the presence and absence of a PRP28/URA3 plasmid. WT = prp28∆ PRP28/URA3 . B) Growth of a prp42∆ strain in the presence of either PRP42 or prp42-101 plasmids. C) Deletion of PRP42 cannot bypass Prp28p. Strains (sectors 3 and 4) containing deletions of both PRP42 and PRP28 and a PRP42/PRP28/URA3 plasmid were constructed and streaked to plates containing 5-FOA to counter-select the PRP42/PRP28/URA3 plasmid. Positive and negative controls are shown in sectors 1 and 2, respectively.
165 A 16˚C 25˚C 30˚C 37˚C
WT
prp42-101 prp28∆ prp42-101
B 16˚C 25˚C 30˚C 37˚C
WT
prp42-101
C 1 2
3 4
166 Figure 3.17. Time course study of splicing in wild-type and prp28∆ prp42-101 extracts. Splicing reactions were incubated for the number of minutes indicated at the top of each lane. The extracts used and temperature of incubation are also noted above the lanes. Diagrams of the splicing products are shown on the right.
167 25˚C 16˚C prp28∆ prp28∆ WT prp42-101 WT prp42-101 5 10 15 20 5 10 15 20 5 10 15 20 5 10 15 20
168 Figure 3.18. prp42-101 is synthetically lethal with bypass alleles of YHC1. A) Genetic scheme for testing alleles of YHC1 for synthetic lethality with prp42-101. A prp42∆ yhc1∆ strain carrying a counter-selectable PRP42/YHC1/URA3 plasmid and a prp42-101 plasmid was transformed with the mutant alleles of YHC1 (yhc1*) and then selected on 5-FOA plates. B) Spot test analysis of strains on media containing 5-FOA. Incubation temperatures are shown at the top of the panels. Bold characters represent the non-bypass version of U1-C.
169 A B 16˚C 25˚C 30˚C - WT prp42::KAN A yhc1::HIS3 PRP42/YHC1 yhc1* C D LEU2 URA3 E prp42-101 F G TRP1 H I K M Plate to 5-FOA N P Q R S T V W Y
170 Figure 3.19. snu71-101 is synthetically lethal with bypass alleles of YHC1. A) Genetic scheme for testing alleles of YHC1 for synthetic lethality with snu71-101. A snu71∆ yhc1∆ strain carrying a counter-selectable SNU71/YHC1/URA3 plasmid and a snu71-101 plasmid was transformed with the mutant alleles of YHC1 (yhc1*) and then selected on 5-FOA plates. B) Spot test analysis of strains on media containing 5-FOA. Incubation temperatures are shown at the top of the panels. Bold characters represent the non-bypass version of U1-C.
171 A B 30˚C 25˚C 16˚C − WT snu71::KAN A yhc1::HIS3 C yhc1* SNU71/YHC1 D E LEU2 URA3 F snu71-101 G H
TRP1 I K M N P Plate to 5-FOA Q R S T V W Y
172 Figure 3.20. Mutations altering a distinct region of Prp8p are synthetically lethal with prp28∆ prp42-101. Features of the Prp8p map are as in figure 1.9. Mutations throughout PRP8 were tested for synthetic lethality with prp28∆ prp42- 101. Alterations that caused synthetic lethality map to an approximately 300 amino acid region of Prp8p. Dotted lines represent the amino acid changes caused by prp8-501 and solid lines represent amino acid alterations that also caused synthetic lethality.
173 1964-1968
1 236 362 611 684 788 861 1094 1197 1371 1608 1817 1861 2065 2413 174
D1778G/R/E E1832A,V W1608R L1794F N1883D M1397I
Q1677F, T1861P D1700E I1825K, L1835F Figure 3.21. C-terminal truncations of SM proteins B, D1 and D3 cannot bypass Prp28p. Strains were streaked to media containing 5-FOA to counter-select a plasmid carrying wild-type PRP28: sector 1 - prp28∆ YHC1-1, sector 2 - prp28∆ SmB∆C/SmD3∆C , sector 3 - prp28∆ SmD3∆C , sector 4 - prp28∆ SmD1∆C , sector 5 - prp28∆ SmB∆C.
175 1 2
63
5 4
Figure 3.22. Deletion of MUD1 cannot bypass Prp28p. Four mud1∆ prp28∆ isolates cannot survive without a plasmid carrying wild-type PRP28 (sectors 3-6). Positive and negative controls are shown in sectors 1 and 2, respectively.
176 CHAPTER 4
DISCUSSION
4.1 Investigation of Alterations in the U1-C Protein and the U1 snRNA that
Allow Bypass of PRP28
4.1.1 Characterization of the YHC1-1 Bypass Suppressor
Altering position L13 of the U1-C protein allows bypass of Prp28p, a normally essential splicing factor proposed to be an RNA helicase. This work provides further characterization of this unique event. As predicted, the cold- sensitiveprp28∆ YHC1-1 strain displayed a temperature-dependent block to splicing in both in vivo and in vitro assays (figures 3.1 and 3.2). The in vitro splicing assay also revealed that the block to splicing at low temperatures occurred before the first catalytic step of splicing, as no intermediate splicing products were observed. This is consistent with the knowledge that Prp28p and the U1-C protein are needed during formation of the pre-spliceosome, disruption
177 of which should block the earliest stages of splicing. The in vitro splicing assay
was further utilized to demonstrate that recombinant His6-Prp28p is able to target and reactivate the stalled splicing complexes in the prp28∆ YHC1-1 extract.
Interestingly, while the recombinant His6-Prp28 protein was able to fully rescue the splicing defect found in prp28-102 extract, splicing in the prp28∆ YHC1-1 extract was not fully rescued to the level of splicing found at permissive temperature (figure 3.4). This could simply be due to variation in extracts or an insufficient level of function of the recombinant His-tagged Prp28 protein.
Another possible interpretation of this result is that the YHC1-1 mutation alone may cause some minor splicing defect that is not sufficient to severely hinder cell growth but that is detectable in my in vitro assays. This possibility is supported by findings that even at “permissive” temperatures the prp28∆ YHC1-1 cells may have some minor splicing defects. Although it is difficult to determine whether these defects detected in the prp28∆ YHC1-1 cells are due to the loss of Prp28p or the mutation in YHC1, there are several interesting observations to consider.
First, splicing in the prp28∆ YHC1-1 extract occurs with slower kinetics than in the wild-type extract at 25˚C, consistent with the slightly slower growth rate observed in the bypass suppressor strain at this temperature (figure 3.3).
177 Second, the RT-PCR results used to confirm the results of the microarray analysis reveal that some unspliced QCR9 transcripts are observed even at the permissive temperature of 30˚C (figure 3.10 A lane 7). Interestingly, while RT-
PCR analysis of two of the candidates confirmed the presence of splicing defects under non-permissive conditions, I was able to determine that the extent of this defect varied between the two samples. In the case of the RPL36B transcript, I observed a drop in mRNA and an increase in pre-mRNA only at 16˚C. For the
QCR9 message, however, a drop in mRNA and increase in pre-mRNA were observed even at the “permissive” temperature of 30˚C. This is an intriguing result, as it indicates unique affects of the prp28∆ YHC1-1 mutations even among the transcripts that were identified by microarray analysis to have splicing defects.
4.1.2 Alterations in the U1 snRNA Bypass PRP28
Mutations in the U1 snRNA that are predicted to disrupt it’s base pairing with the 5’ SS and thus destabilize the U1/5’ SS interaction were also found to bypass PRP28. This result provided a valuable clue as to how alteration of the
U1-C protein may be allowing bypass and has additional significant implications
178 on the general understanding of RNA helicase function. Comparable to the duplex-destabilizing U1 snRNA , it is likely that alteration of the U1-C protein also weakens the U1/5’ SS interaction, thus eliminating the requirement for Prp28p to help in the disruption of the U1 snRNA/5’ SS duplex (figure 3.7). This model is consistent with the finding that bypass of PRP28 is no longer feasible at low temperature. Under these conditions, the U1/5’ SS interaction would be thermodynamically re-stabilized, and PRP28 would again be required to help disrupt the complex. Our model that changes to U1-C destabilize the U1/5’ SS duplex was further supported by data showing that the L13S alteration could suppress in vivo splicing defects due to an extension of the U1 snRNA/5’ SS base pairing (Chen et al., 2001). Importantly, our findings have contributed to the understanding of how DExD/H-box proteins may function. Our discovery that
Prp28p, a putative RNA helicase, can be bypassed by altering U1-C, a protein factor, indicates that RNAs are not the only molecules impacted by Prp28p.
Instead, it is possible that Prp28p may target a more complex structure that includes both RNA and proteins and may help rearrange the interactions among them, leading to the release of the U1 snRNP from the pre-mRNA. The theory of
RNA helicases functioning at RNPases is rapidly gaining support, as a number of
179 other laboratories have recently come to similar conclusions. In the splicing field, it was found that the deletion of MUD2, a gene encoding a protein that binds near the branch point, can bypass the requirement for another putative DExD/H box helicase, Sub2p (Kistler and Guthrie, 2001). The deletion of another splicing gene, CUS2, can bypass the requirement for ATP during U2 binding to the branch point. The ATPase requirement is likely attributed to the DExD/H box protein Prp5p and that Cus2p brokers the interaction between Prp5p and the U2 snRNP (Perriman and Ares, 2000). Finally, the vaccinia virus NPHII RNA helicase was shown to displace a protein bound to an RNA substrate in an in vitro unwinding assay, directly showing that an RNA helicase can rearrange protein/RNA contacts. Therefore our laboratory’s finding that altering the U1-C protein can bypass Prp28p is a strong indicator that Prp28p may normally aid in the rearrangement of both RNA and proteins, leading to U1 snRNP release.
4.1.3 Growth Phenotypes of the U1-C L13 Variants
Our working model indicates that position L13 within the U1-C protein is likely to be involved in an intermolecular interaction that normally helps to stabilize the association between the U1 snRNP and the 5’ SS. Despite much
180 support for this idea, we have not yet identified the factor(s) that this region of
U1-C may be contacting. The initial finding that the group of amino acids that could not bypass were all hydrophobic in nature indicated that position L13 was likely to be involved in an interaction that requires a hydrophobic residue at this position. We hypothesized that certain amino acid changes may cause a greater disruption than others and therefore provide a more detailed picture of this putative interaction. For example, amino acids with highly charged or extremely large side chains might cause more severe disruptions than those with side chains more similar to that of leucine. I first tested whether any of the L13 variants were more or less suitable as substitutes for the normal U1-C protein in an otherwise wild-type cell. Surprisingly, only minor growth defects were observed in a few mutants at high or low temperatures, indicating that all of these alleles were able to function at or near the same level as wild-type (figure 3.8).
This result is intriguing; despite the strong evolutionary conservation of position
L13, changing this residue to any other amino acid appears to have little impact on the function of the protein, even at less than optimal temperatures. The fact that position L13 is strictly conserved among species indicates that there is a reason for maintaining a leucine at that location. My findings, however, argue
181 otherwise, as any amino acid substitution at this position not only leads to a viable cell but does not significantly impact growth, even at extreme temperatures. We were also hopeful that different amino acid substitutions at position 13 would allow bypass of PRP28 to different extents to provide clues as to the nature of the interactions that this residue participates in. Unexpectedly, however, all of the bypass alleles demonstrated a very similar phenotype. While this may not provide us with the information we were initially seeking, it may indicate that this interaction could function as an on/off switch. In other words, if a sufficient interaction takes place, splicing can proceed without major difficulties.
If the interaction is not sufficient, regardless of what amino acid change is causing the disruption, splicing is impeded. Although the L13 alterations resulted in very similar bypass phenotypes, it is worth noting that they are all cold- sensitive and temperature-sensitive, a result that supports our model of how bypass occurs. At low temperatures the U1/5’ SS duplex may become thermodynamically stabilized, preventing bypass. At high temperatures, the complex may be too unstable or not form at all, resulting in cell lethality. It is also important to note that the apparent lack of growth phenotypes in the strains containing the various L13 alterations may not closely reflect the effects they
182 have on splicing. As seen above with the in vivo and in vitro splicing assays, more sensitive in vitro analysis might be required to detect slight differences among these variants. Therefore it will be of interest to utilize microarray techniques to analyze the L13 variants, as this may provide a more complete picture of their impacts on splicing.
4.1.4 A Novel Role for U1-C at the 5’ SS
A recently published report by Du and Rosbash (2002) has suggested a new model for 5’ SS recognition. This model is based on their finding that the
U1-C protein is able to recognize and bind to the nucleotide sequence GUAU, a portion of the 5’ SS that also base pairs with the U1 snRNA. The authors also found that while the U1-C/pre-mRNA interaction occurs at low temperature, the
U1 snRNA/pre-mRNA interaction does not. When U1-C was depleted, however, the RNA-RNA duplex was able to form, indicating that the U1-C protein suppresses RNA base pairing at low temperatures. This led to their hypothesis that the U1-C protein may be the first factor to identify and bind to the 5’ SS, subsequently giving way to the interaction between the U1 snRNA and the 5’ SS.
Although there is still no direct evidence for this model, it is also supported by its
183 similarity to the process of branch point recognition, where protein factors clearly identify this region before the U2 snRNA makes contact. Furthermore, the authors suggest that the mutations altering position L13 within U1-C may allow spliceosome assembly to proceed without U1 snRNA/5’ SS base pairing, an idea that is consistent with our model that Prp28p is bypassed by weakening this interaction.
This model also raises the possibility that Prp28p may play an additional role in removing U1 snRNP particles that erroneously bind to GUAU sequences that are not authentic splice sites. The sequence GUAU is likely to occur multiple times in any given transcript, providing a number of recognition sites for the U1-C protein. The observation that more than one U1 particle appears to bind to a single pre-mRNA supports this idea (Eperon et al., 1993). After the U1 particles bind to the pre-mRNA through the U1-C protein, however, the association may not be productive if the additional sequences required for U1 snRNA binding are not present. In this case, Prp28p may help to remove these dead-end complexes, acting to sweep the transcript clean to allow splicing to proceed. It is worth noting that Staley and Guthrie (1999) observed marked degradation of actin transcripts containing hyperstabilized U1/5’ SS duplexes in a prp28-1
184 mutant strain, prompting them to propose that there may be a discard pathway for transcripts with aberrant 5’ splice sites. This model may also apply to transcripts containing multiple U1 snRNP particles bound to 5’ SS-like GUAU sequences that cannot be removed due to mutations in PRP28.
4.2 Novel Mutations in PRP42 and SNU71 Can Bypass PRP28
4.2.1 prp42-101 and snu71-101 May Destabilize the U1/5’ SS Duplex
The novel alleles prp42-101 and snu71-101 were identified as bypass suppressors of PRP28, genetic interactions indicating that Prp42p and Snu71p may normally play a role in stabilizing the U1 snRNP/5’ SS association. This prediction is based on our previous finding that Prp28p can be bypassed by alterations of the U1-C protein, a factor known to bind to the 5’ SS, as well as U1 snRNA mutations predicted to diminish binding to the 5’ SS (Chen et al., 2001;
Du and Rosbash, 2002). Consistent with the phenotypes caused by the YHC1 bypass alleles, both prp42-101 and snu71-101 cause cold-sensitive growth phenotypes in the absence of PRP28 (figure 3.16 and R. Hage, unpublished).
This conditional phenotype may indicate that at low temperatures the U1/5’ SS interaction that is weakened by these mutations is thermodynamically re-
185 stabilized, thus reinstating the requirement for PRP28. In addition, prp42-101 and snu71-101 are synthetically lethal with the YHC1 bypass alleles, further supporting the hypothesis that these mutations have similar effects during splicing. Combining two mutations that each relax the U1/5’ SS duplex, even slightly, may result in a U1 particle that binds to pre-mRNAs extremely weakly or possibly not at all, culminating in cell inviability. Interestingly, there were a few exceptions to this trend. In the case of prp42-101, the mutation causing the
L13M change in U1-C caused synthetic lethality, even though this allele does not allow bypass of PRP28. Similarly, the non-bypass mutation causing the L13Q alteration in U1-C was synthetically sick with snu71-101. Although the meaning of these inconsistencies is currently unclear, they do indicate that the interactions allowing bypass with each of these factors is unique and may provide important clues into the intricacies of their bypass mechanisms.
4.2.2 prp42-101 is Synthetically Lethal with Alleles of PRP8
Further support for the idea that the additional bypass suppressors are involved in the same control circuit as U1-C is provided by the determination that prp42-101 demonstrates synthetic lethality with a number of PRP8 alleles in a
186 pattern similar to that found for YHC1-1. This finding likely reflects the general interaction between Prp8p and Prp28p and does not necessarily indicate that
Prp8p is directly interacting with both U1-C and Prp42p. In other words, the same region of Prp8p becomes critical for regulating spliceosomal rearrangements regardless of why Prp28p is absent. Although the same general region of Prp8p was found to be critical in the presence of both the YHC1-1 and prp42-101 alleles, the patterns were slightly different. For example, prp8-124 was synthetically lethal with prp28∆ prp42-101 but not with prp28∆ YHC1-1. This allele causes an alteration at amino acid 1397, a position approximately 200 amino acids out side the proposed regulation domain. This may indicate a slight difference in these two interactions. In addition, the allele causing the D1670N and D1778G alterations may display dominant sythetic-lethality with prp28∆ prp42-101, as no viable transformants were obtained with this allele. Although this result is not conclusive, it also demonstrates the uniqueness of each bypass allele. Finally, it will be of interest to determine whether alterations in the same region of Prp8p also interact with the additional bypass alleles of SNU71 and
YNL187W.
187 4.2.3 Protein-Protein Interaction Domains May be Affected by prp42-101 and snu71-101
Although we initially predicted that Prp28p might be bypassed by altering proteins known to localize near the 5’ SS, Prp42p and Snu71p are not among the protein factors found to cross link to this region of the pre-mRNA (Zhang and
Rosbash, 1999). The presence of potential protein-protein interaction motifs in both of these factors, however, may provide clues as to how they may contribute to the stabilization of the U1/5’ SS association despite their apparent lack of proximity. It seems likely that these two proteins may have a more indirect influence on the U1/5’ SS interaction than does the U1-C protein. One possibility is that through protein-protein interactions Prp42p and Snu71p may help to stabilize the U1-C/pre-mRNA interaction. Alternatively, it is possible that their protein interaction domains are important for maintaining contacts crucial to the overall structure of the U1 snRNP particle. Alterations disrupting these contacts may compromise the general framework of the U1 snRNP, leading to a weaker interaction with the pre-mRNA and therefore bypass of Prp28p. This model may be supported by the finding that the U1 snRNP appears to adopt an altered configuration in extracts depleted of Prp42p (McLean and Rymond, 1998).
188 Although the precise mechanism by which these factors allow bypass of
Prp28p is currently unclear, it is apparent that disruptions to the protein interaction domains are likely the key. The prp42-101 mutation is predicted to alter an amino acid in one of the conserved TPR repeats, a change that is likely to disrupt an important protein-protein interaction. Alternatively, this change could alter an intramolecular TPR-TPR interaction within Prp42p, leading to a conformational defect that could in turn disrupt the protein’s function. The snu71-
101 mutation falls between two conserved SR-like dipeptides. Although the effect of this mutation remains to be determined, it is possible that these motifs may be somehow compromised in the resulting protein.
4.2.4 Prp42p May be Required for the Efficient Splicing of a Subset of
Transcripts
Although the alterations in Prp42p and U1-C appear to allow bypass of
Prp28p by a similar mechanism, the difference between the prp42-101 and the
YHC1-1 bypass alleles is apparent in their in vitro splicing phenotypes. Although both bypass strains are cold-sensitive in the absence of PRP28, only the prp28∆
YHC1-1 combination results in a clear cold-sensitive in vitro splicing defect.
189 Splicing does appear to be slowed in the prp28∆ prp42-101 extracts, but clearly occurs even at low temperatures (figure 3.17 and data not shown). This could simply be due to an overall weaker influence of the prp42-101 allele that once placed in the altered context of the in vitro assay is not able to produce a striking phenotype. An alternative explanation is that Prp42p may not play a direct role in the splicing of all cellular pre-mRNAs. My in vitro assay only tested the splicing of the actin pre-mRNA, and it is possible that the altered Prp42p does not have a severe impact on this particular message. As an essential splicing factor, however, Prp42p is most likely required for the efficient splicing of a subset of transcripts. In this scenario, although not all messages would be affected by mutations in PRP42, a severe splicing defect in even one critical transcript could prevent cell growth. Previous studies provide strong support for this model.
After being identified as synthetically lethal with mutations in the U1 snRNA, an allele of prp42 (mud16) was tested for splicing defects. The only deficiencies observed were minor splicing defects in transcripts that were known to be inefficiently spliced or that harbored a mutant 5’ SS sequence (Gottschalk et al.,
1998). There is also precedence for an essential, intron-specific splicing factor that is part of the U1 snRNP. A phenomenon similar to what I have found with
190 PRP42 was observed during the study of another U1 snRNP factor, LUC7.
Luc7p appears to bridge the interaction between the cap binding complex (CBC) and the U1 snRNP and was shown to promote cap-proximal 5’ SS selection. A luc7-1 strain is temperature sensitive, but primer extension assays show that the in vivo splicing of wild-type transcripts was largely unaffected at the non- permissive temperature. Reporter transcripts containing mutant 5’ SS or branchpoint sequences, however, were inefficiently spliced (Fortes et al., 1999a).
Intriguingly, there appears to be a connection between LUC7, PRP42 and
SNU71 that may, at least in part, account for the similarities noted above. First, a mutation in SNU71 (LUC5) was identified in the same screen as LUC7 in a search for factors that interact with the CBC (Fortes et al., 1999b). This is not unexpected, as cooperative interactions between the CBC and splicing factors have previously been identified (Lewis et al., 1996a; Lewis et al., 1996b).
Significantly, both Prp42p and Snu71p are absent from U1 snRNPs purified from luc7-1 mutant extracts (Fortes et al., 1999a). Therefore in the luc7-1 cells where splicing appeared largely unaffected at the non-permissive temperature, Prp42p and Snu71p were most likely also absent from the U1 snRNPs. This observation
191 led the authors to suggest that some of the U1 snRNP proteins including Luc7p,
Snu71p and Prp42p may only be required for the splicing of certain introns.
If it is true that Prp42p and Snu71p are not required for the splicing of all transcripts, what might their actual role be? Our data implicates these factors in
5’ SS selection, an idea that is corroborated by the observation that while wild- type transcripts were clearly spliced in mud16 (prp42) mutant cells, messages containing an altered 5’ SS were not (Gottschalk et al., 1998). Additional support for these predicted functions may be revealed by a comparison of the yeast and mammalian U1 snRNP particles and their roles in splice site selection. An unanswered question in the field of splicing is the reason for the apparent lack of homology between the yeast and mammalian U1 snRNP particles. The yeast U1 snRNP contains at least 17 specific proteins, of which only three are found in its mammalian counterpart. Concurrently, only metazoan systems contain the large
SR family of proteins that are involved in 5’ SS selection and allow U1 snRNP- independent splicing when present in high concentrations (Crispino et al., 1994;
Tarn and Steitz, 1994). Although a defined family of SR proteins does not exist in yeast, there are a number of factors with similar SR motifs, including Snu71p.
It has therefore been suggested that some of the yeast U1 snRNP proteins may
192 be functionally homologous to the mammalian SR factors (Gottschalk et al.,
1998; McLean and Rymond, 1998). A possible explanation for this difference in organization lies in the fact that the splicing of mammalian transcripts is much more complex than that of yeast transcripts. While few yeast genes contain introns, most mammalian transcripts have numerous introns that may also undergo regulated alternative splicing. This complex regulation is mediated in part by variations in splice site sequences in combination with the SR proteins that help identify the proper splice sites under the appropriate conditions. In yeast, however, there are many fewer introns and the splice site sequences are more highly conserved. Therefore it is possible that in yeast the factors that aid in splice site recognition are integral components of the U1 snRNP while the mammalian factors only transiently associate during the splicing of certain transcripts (Fabrizio et al., 1994; Gottschalk et al., 1998).
This idea correlates with the hypothesis that some factors including
Prp42p, Snu71p and Luc7p may only be required for the splicing of certain transcripts. It also raises the possibility that under true in vivo conditions, all of these proteins may not always be present in every U1 particle, a model that might explain the observed weak association of some of the factors including
193 Prp42p. The idea that these yeast proteins are functionally similar to the mammalian SR proteins is also supported by findings that these yeast factors appear to aid in the recruitment of other proteins to the transcript, much as the
SR proteins do. In particular, Snu71p and Luc7p have been shown to interact with the CBC. These interactions alone implicate them in 5’ SS selection, as the
CBC is known to increase cap-proximal 5’ SS recognition by the U1 snRNP
(Colot et al., 1996; Lewis et al., 1996a; Lewis et al., 1996b). Interestingly, homologues of Luc7p in several more complex organisms including humans, C. elegans and Drosophila all contain carboxy-terminal extensions with numerous
SR and SR-like repeats (Fortes et al., 1999a). Naturally, there are alternative models that must be considered. For example, it is possible that mammalian homologues to some of these yeast U1 proteins do exist, but that they have not yet been identified as they are less tightly associated and lost during biochemical fractionations (Gottschalk et al., 1998).
194 4.3 Bypass of PRP28 is a Specific Event
4.3.1 Tail-truncated Versions of Three Sm Proteins Cannot Bypass PRP28
I identified Sm proteins B, D1 and D3 as potential components of the target of Prp28p based on the reports that these proteins, like U1-C, contribute to
U1 snRNP/5’ SS complex stability and are physically adjacent to U1-C (Stark et al., 2001; Zhang et al., 2001). My finding that the tail-truncated Sm mutants that weaken commitment complex formation could not bypass Prp28p indicates that these three proteins are unlikely to be part of Prp28p’s target. One possible explanation for this may be the positioning of these Sm proteins within the U1 snRNP/5’ SS complex. Although the three-dimensional structure of the human
U1 snRNP indicates that the U1-C protein is positioned above the Sm proteins and is therefore physically removed from the pre-mRNA, other data shows that the U1-C protein is in direct contact with the pre-mRNA. In addition to the known cross-links between U1-C and the pre-mRNA, the U1-C protein directly contacts the 5’ SS by binding to the conserved GUAU sequence (Du and Rosbash, 2002;
Zhang and Rosbash, 1999). In light of this information, my initial hypothesis that
U1-C may be helping to position the Sm proteins from its location above them seems unlikely. Although there could simply be differences between the yeast
195 and human particles, the apparent contradiction between the structural findings and the U1-C/5’ SS interaction data could also be explained by conformational changes that may occur within the U1 snRNP. The U1 particles purified for structural analysis were not bound to pre-mRNAs and therefore may not have been in their “active” state. It is likely that U1 snRNPs can assume multiple conformations, including one that is adopted during the association with the pre- mRNA and another that does not allow the particle to bind to transcripts. In this scenario, U1 snRNPs would undergo various structural rearrangements that allow binding to and then release from transcripts. These multiple conformations could account for the observed contacts between numerous U1 proteins and the pre-mRNA as well as the apparent three-dimensional structure.
Regardless of whether any structural rearrangements occur within to U1 snRNP during binding to the 5’ SS, cross-linking data indicate that the positions of the Sm protein contacts with the pre-mRNA may simply be too distant from
Prp28p’s target. Specifically, both SmD1 and SmD3 cross-link to position –2 in the exon while SmB cross-links most strongly to position +2 and only weakly to position +6 in the intron. The U1-C protein was found to strongly cross-link only to intron position +6 (Zhang and Rosbash, 1999). This indicates that although
196 the Sm proteins do contact the pre-mRNA to stabilize the commitment complex, they may simply be too distant from Prp28p’s proposed target site, the U1-C/pre- mRNA contact at position +6. The weak cross link of SmB to position +6 is likely a result of its physical contact with U1-C (Nelissen et al., 1994). Despite being the closest candidate, the tail truncated SmB protein could not bypass Prp28p.
This is not completely unexpected, however, as the tail truncation of SmB alone was found to have only a small impact on commitment complex formation. A severe defect was not detected until the SmB and SmD3 tail truncations were combined (Zhang et al., 2001). In my system, however, the SmB and SmD3 double mutant was still unable to bypass Prp28p.
4.3.2 Deletion of MUD1 Cannot Bypass PRP28
Although we felt it was unlikely that Mud1p and Nam8p would be included in the target of Prp28p, it remained a possibility that any significant disruption to the U1 snRNP could affect the particle’s ability to bind to transcripts and thus allow bypass Prp28p. We therefore decided to ask whether the deletion of these non-essential components of the U1 snRNP could bypass PRP28. Our finding that PRP28 cannot be bypassed by the deletion of MUD1 or NAM8 or even by
197 the Sm tail truncations known to weaken the U1/5’ SS interaction indicates that the bypass suppressors we have identified are likely to be true and specific components of the target of this putative RNPase. Prp28p cannot be bypassed by any kind of mutation that disrupts the U1 snRNP and/or destabilizes its association with the 5’ SS. Therefore it appears that Prp28p specifically interacts with distinct components of the commitment complex and does not randomly counteract any protein that happens to be part of the stabilizing structure.
Importantly, my finding that Prp28p is unlikely to be responsible for destabilizing the interaction between the Sm proteins and the pre-mRNA indicates that additional factors are likely crucial in aiding in the displacement of these components. One possible candidate is Dbp2p, another DEAD-box protein. The human homologue of Dbp2p, p68, has been implicated in mediating the U1/5’ SS interaction (Liu, 2002). Although this data does not discount the role of Prp28p at this stage in splicing, it indicates that additional factors may also aid in rearranging these interactions.
198 4.4 The Function of Prp8p at the Catalytic Core
4.4.1 A Specific Domain Within Prp8p Appears to Aid in the Removal of U1
From the 5’ SS
A screen designed to identify splicing factors that assist Prp28p in removing the U1 snRNP from the 5’ SS revealed a novel mutation in the gene
PRP8. This finding is consistent with previous data showing that Prp8p is positioned at the catalytic core of the spliceosome and may regulate spliceosome activation. Although there are no obvious domains or motifs within Prp8p, several potential functional regions within the protein have been identified. My data indicate that a specific region of Prp8p may be required to assist in the disruption of the U1/5’ SS interaction. This putative domain lies very near the portion of Prp8p that was found to cross-link to the 5’ SS (Reyes et al. 1999). It also partially overlaps the regions of Prp8p that have been implicated in pyrimidine tract recognition, 3’ SS fidelity, suppression of splice site mutations, and suppression of a mutation in the U4 snRNA (figure 3.12) (Collins and
Guthrie, 1999; Kuhn and Brow, 2000; Siatecka et al., 1999; Umen and Guthrie,
1996). As might be predicted, none of the mutations previously found to cause defects in pyrimidine tract recognition were found to be synthetically lethal with
199 prp28∆ YHC1-1 or prp28∆ prp42-101. The defects that disrupt Prp8p’s function
in pyrimidine tract recognition are unlikely to also affect its function at the 5’ SS
as it was shown that the domains governing pyrimidine tract recognition and
splice site suppression were clearly separable (Collins and Guthrie, 1999). I also
found that the prp8-L280P mutation that can suppress prp28-1 is not
synthetically lethal with our bypass mutants (Kuhn et al., 2002). This finding is
consistent with the fact that as a suppressor, this version of Prp8p is somehow
helping to compensate for a defect in Prp28p’s normal function, making it unlikely
to cause synthetic lethality in Prp28p’s absence.
A general trend in the results of this analysis is that mutations originally
identified as suppressors of splice site mutations were not synthetically lethal
with our U1-C bypass mutant, while those that were synthetically lethal were
unable to suppress splice site mutations. This may be explained by the finding
that the suppression of the splice site mutants by specific alleles of PRP8 was
attributed to an enhancement of the second step of splicing (Collins and Guthrie,
1999; Kuhn and Brow, 2000; Siatecka et al., 1999; Umen and Guthrie, 1996). In
contrast, alterations in PRP8 that cause synthetic lethality with the prp28∆YHC1-
1 or prp28∆ prp42-101 bypass mutants are likely to be affecting an earlier time
200 point in spliceosome activation, specifically before the first catalytic step. There are, however, a few interesting exceptions to this trend. The first is prp8-201, the allele initially identified as a suppressor of U4-cs1 (Kuhn et al., 1999). This allele was synthetically lethal with both bypass mutants, and unlike any of the other U4- cs1 suppressor mutations that were tested, was also found to suppress splicing defects due to 5’ SS and 3’ SS mutations (Kuhn and Brow, 2000). As has been previously noted, prp8-201 appears to be a rather unique allele that results in multiple phenotypes including temperature sensitivity, indicating that the interaction detected in my study is not necessarily linked to its splice site suppressor activity. The genetic interaction between our bypass mutants and prp8-201 is also unlikely to be due to its U4-cs1 suppressor phenotype as none of the alleles tested that were identified in a secondary screen for U4-cs1 suppressors showed synthetic lethality with prp28∆ YHC1-1 (Kuhn and Brow,
2000).
The second exceptions to the aforementioned trend are prp8-122 and prp8-124, two of the alleles initially found to suppress a 3’ SS mutation and later re-classified as a general splice site suppressors (Collins and Guthrie, 1999;
Umen and Guthrie, 1996). Both alleles were synthetically lethal with prp28∆
201 prp42-101 and prp8-122 was also synthetically lethal with prp28∆ YHC1-1.
Interestingly, these alleles and prp8-201 were also found to exacerbate a hyper- stabilized U1/5’ SS duplex (table 3.1) (C.Collins, personal communication, Kuhn and Brow, 2000). Therefore, while both prp8-201, prp8-122, and prp8-124 are splice site suppressors, they are also able to exacerbate a hyper-stabilized U1/5’
SS duplex, indicating that this may be the effect of these mutations that is likely to explain the observed interactions with our bypass mutants.
4.4.2 A Role for Prp8p in Spliceosomal Activation
My results add to a growing body of evidence indicating that Prp8p is positioned at the catalytic core of the spliceosome and acts to regulate and coordinate the various rearrangements that are necessary to activate the spliceosome. How might this coordination be achieved? Prp8p interacts with both splice sites as well as the U5 snRNA, interactions that implicate this factor in juxtaposing the 5’ and 3’ splice sites during catalysis (Dix et al., 1998;
Teigelkamp et al., 1995). Additional interactions with the U6 snRNA also indicate a role in delivering this putative catalytic element to the core, suggesting that
Prp8p plays a direct role in positioning the critical players for catalysis (Vidal et
202 al., 1999). Furthermore, Prp8p appears to play a direct role in regulating the disruption of the U4/U6 interaction and coupling this to the displacement of U1 from the 5’ SS, rearrangements that are linked and necessary for spliceosomal activation (Cheng and Abelson, 1987; Konarska and Sharp, 1987; Konforti et al.,
1993; Kuhn et al., 1999; Madhani and Guthrie, 1992; Staley and Guthrie, 1999).
Prp8p’s participation in these rearrangements is supported by a large amount of data.
First, Prp8p demonstrates numerous interactions with the 5’ SS region.
The C-terminal region of the human homologue of Prp8p can be cross-linked to the conserved GU of the 5’ SS (Reyes et al., 1996; Teigelkamp et al., 1995;
Wyatt et al., 1992). Prp8p also participates in interactions with several factors known to associate with the 5’ SS. Genetic interactions have been found between PRP8 and PRP28 (Kuhn et al., 2002; Strauss and Guthrie, 1991).
Physical interactions have been observed between Prp8p and the U1 snRNP proteins Prp40p, Prp39p and Snp1p (Abovich and Rosbash, 1997; van Nues and
Beggs, 2001). Interestingly, Snp1p retrieved Brr2 in an exhaustive 2-hybrid screen (Fromont-Racine et al., 1997), indicating one possible means of communication between the U1 and U4/U6/U5 snRNPs. Another sign of the
203 connection between the U1 and U5 snRNPs was an observed interaction between the 5’ end of the U1 snRNA that contacts the 5’ SS and the U5 snRNA
(Ast and Weiner, 1997).
Second, several lines of evidence also demonstrate that Prp8p influences and may regulate U4/U6 unwinding. As a component of the U5 snRNP, Prp8p is affiliated with U4 and U6 in the tri-snRNP. Furthermore, Prp8p was found to cross link to the U6 snRNA at position U-54. This provides a physical link between Prp8p and the region of U6 that is directly adjacent to both the ACAGA box that binds to the 5’ SS and stem I of the U4/U6 duplex. It is this duplex that is likely unwound by Brr2p during spliceosome activation and that contains the region of U6 that subsequently pairs with the U2 snRNA and has been implicated in catalysis (Vidal et al., 1999). Mutations in PRP8 were also found to suppress the U4-cs1 mutation that hyper-stabilizes U4/U6 stem I and thus blocks U4/U6 unwinding (Kuhn and Brow, 2000; Kuhn et al., 1999). A direct link between
Prp8p and Brr2p was established when both were identified in a screen for factors that were synthetically lethal with a mutation in the U2 snRNA (Xu et al.,
1998). Prp8p is also involved in two separate and mutually exclusive interactions with Brr2p during splicing, indicating that a conformational change in Prp8p may
204 take place during spliceosome activation and that a resulting change in interactions with Brr2p may be a means of regulating the helicase (van Nues and
Beggs, 2001).
It has also been proposed that Prp8p may mediate the functions of both
Prp28p and Brr2p as a way to regulate spliceosome activation (Kuhn et al.,
2002). The authors of this study suggest that Prp8p may help coordinate the
U1/5’ SS and U4/U6 rearrangements by partially masking the substrates of these putative helicases until spliceosomal rearrangements are needed. Specifically, genetic interactions implicate the N-terminal region of Prp8p in Prp28p regulation and a more C-terminal region in Brr2p regulation. Although the model that an N- terminal region of Prp8p may be regulating Prp28p may seem to be at odds with my data showing that a more C-terminal region of Prp8p may be assisting
Prp28p in displacing U1 from the 5’ SS, these findings are not necessarily contradictory. It is possible that these two regions of Prp8p actually lie close to one another as a predicted intra-molecular fold within Prp8p would bring the
Brr2p regulation domain near our new U1 displacement domain (see figure 3.12)
(Kuhn et al., 2002). It would be interesting to determine whether this is the same region of Prp8p that contacts the U6 snRNA. If this were true, it might indicate
205 that this region of Prp8p may be initially masking the U4/U6 duplex to inhibit
Brr2p and subsequently switch to guiding the U6 snRNA to the 5’ SS during activation.
Another possibility is that Prp8p has two separate functions that affect
Prp28p. The N-terminal region may mask part of Prp28p's substrate or otherwise regulate its functions, while a separate region towards the C-terminus may function to help displace U1 from the 5' SS. Interestingly, Prp8p has been reported to interact with U1-C (van Nues and Beggs, 2001). Although the details of this interaction are unknown, it could indicate one means by which Prp8p may block Prp28p’s target. It is possible that Prp8p may initially interact with U1-C, preventing Prp28p from recognizing this part of its substrate. Ensuing conformational changes during may result in Prp8p releasing its contact with U1-
C, thus allowing Prp28p to recognize its target. Alternatively, Prp8p may contact
U1-C in a manner that helps to remove it from the 5’ SS during spliceosome rearrangements. Experiments addressing the timing of this interaction could shed light on the nature of the Prp8p/U1-C relationship.
In addition to Prp8p’s possible role in coordinating Prp28p and Brr2p, may play a very direct role in 5’ SS selection. Maroney et al. (2000) found that Prp8p
206 contacts the 5’ SS independent of the U2 snRNP binding to the branchpoint.
This association requires ATP and incubation at 30˚C and occurs when Prp8p is a part of the tri-snRNP. Based on other reports revealing that U1 snRNP particles bind to many “pseudo” 5’ SS sequences, the authors propose that the tri-snRNP subsequently binds to only authentic splice sites with the help of Prp8p
(Eperon et al., 1993). It is further suggested that the ATP-dependence of this interaction may be attributable to the function of Brr2p, as mutations in BRR2 that debilitate it’s ATPase activity prevent tri-snRNP association with the pre-mRNA
(Raghunathan and Guthrie, 1998). This information suggests that the tri-snRNPs that bind to the 5’ SS may assume a specific conformation, possibly one where
Brr2p has partially dissociated the U4 and U6 snRNAs (Maroney et al., 2000).
This idea is also consistent with the finding of van Nues and Beggs (2001) that two separate interactions occur between Prp8p and Brr2p.
The following working model demonstrates how the factors discussed within this work may function in the switch of U1 for U6 at the 5' SS (figure 4.1).
As the spliceosome first begins to form, the interaction between the U1 snRNP and the 5’ SS is established and stabilized with help from the U1 snRNA, the U1-
C protein and likely additional factors including some of the Sm proteins, Prp42p,
207 and Snu71p (Zhang et al., 2001). As recent evidence indicates that the U1-C protein recognizes and binds to the nucleotide sequence GUAU, it is possible that the U1 snRNP may bind to multiple 5’ SS-like sites in the pre-mRNA after which the true splice site is selected by binding of the tri-snRNP. Prp8p may play a crucial role in the tri-snRNP’s recognition of authentic 5’ SS sequences by interacting directly with the pre-mRNA as well as several U1 proteins including
Prp39p, Prp40p and Snp1p. During these early stages of spliceosome formation
Prp8p also makes contacts with many additional splicing components in order to assure proper assembly and regulate some spliceosomal rearrangements.
These include the 5’ and 3’ splice sites, the U6 snRNA and numerous proteins, possibly including Prp28p and Brr2p. In it’s initial “pre-catalysis” conformation,
Prp8p may also mask all or part of the U1/5’ SS and U4/U6 duplexes as well as possible protein components that are targeted by Prp28p and Brr2p. Based on the proposed intra-molecular folds within Prp8p, it is feasible that the putative
Brr2p regulation domain and the domain identified in this study are near one another and may be masking the U4/U6 duplex. In addition, this may also encompass the region of Prp8p that contacts the U6 snRNA. When spliceosome activation occurs, a conformational change within Prp8p may reveal the
208 substrates of the helicases and/or re-position these proteins in order to activate their functions. At this time Brr2p can resolve the U4/U6 duplex, and Prp8p may help guide the U6 snRNA towards the 5’ SS to compete with the U1 snRNA for binding, thus aiding in the displacement of the U1 snRNP. This idea is supported not only by the cross-link between Prp8p and position U54 of the U6 snRNA but also the observation that the U6 snRNA can indeed compete with the U1 snRNA for binding at the 5’ SS (Staley and Guthrie, 1999; Vidal et al., 1999).
Concurrently, Prp28p could work to resolve the U1 snRNA/5’ SS interaction and counteract the stabilizing effects of U1-C and other proteins including Prp42p and
Snu71p.
4.5 Prospectus
The work presented in this dissertation provides a continuing analysis of the interaction between the U1 snRNP and the 5’ SS. We have identified
Prp42p, Snu71p and Ynl187wp as important factors in this interaction, and it is of great interest to delineate the interactions among these factors in order to further understand their roles in U1/5’ SS regulation. Alterations in Prp42p and Snu71p that allow bypass of Prp28p may disrupt protein-protein interaction domains.
209 This may guide future experiments to determine which proteins within the U1 snRNP are in contact with one another and in turn, which of these may be disrupted as a result of the bypass alterations. Specifically, Prp42p and Snu71p contacts with Luc7p, U1-C and even each other might be predicted. Luc7p has two zinc fingers that warrant particularly close attention as they may mediate important protein-protein contacts (Fortes et al., 1999a). If interactions between
Luc7p and Prp42p and Snu71p are found as expected, it may be possible to isolate mutations in LUC7 that are also capable of bypassing PRP28. After identifying the protein-protein contacts that normally occur, it will then be critical to determine how these contacts are altered in the presence of the bypass mutations. Equally important is the testing of whether any of these factors may directly contact the U1 snRNA or the pre-mRNA. Finally, it will be very interesting to determine the affects these alterations have on the structure of the
U1 snRNP as well as its ability to bind to the 5’ SS during commitment complex formation.
Further analysis of the prp42-101 and snu71-101 mutations is also important to determine whether these mutations affect all transcripts or are delegated to the regulation of 5’ SS recognition of a certain group of transcripts.
210 Future work is likely to include the testing the splicing of transcripts containing various mutations in prp42-101 and snu71-101 extracts, as this may lead to a clear defect and a better idea of the roles of these factors during splicing.
Another method that may allow us to discern whether there is a subset of transcripts that are severely affected by these mutations is the use of splicing microarrays (Clark et al., 2002; Spingola et al., 1999). Preliminary splicing microarray experiments to analyze global splicing defects in the prp28∆ prp42-
101 and prp28∆ snu71-101 strains revealed that there may be subsets of transcripts that are affected in each strain (T.-H. Chang, unpublished). Additional experiments will likely include microarray analysis of strains harboring the prp42-
101 and snu71-101 mutations alone. The resulting data should be valuable in determining the specific defects caused by these alleles and could potentially reveal a specific class of introns that rely on these factors. We are also hopeful that microarray analysis could be utilized to better characterize the defects caused by the YHC1-1 bypass mutation. Specifically, my RT-PCR results indicate that some transcripts may be affected even at “permissive”
211 temperatures. A larger number of transcripts will need to be analyzed with a more quantitative assay such as real-time PCR in order to determine whether this observation may be more meaningful.
Finally, I have also identified a potential region of PRP8 that appears to be important for assisting Prp28p in disrupting the U1/5’ SS interaction. This finding adds to an increasingly detailed picture of the complex series of events that are responsible for spliceosome activation. We are hopeful that our unique prp28∆YHC1-1 prp8-501 strain may allow us to uncouple the spliceosomal rearrangements, allowing us to isolate spliceosomes that are in the process of becoming activated. Such intermediates may be re-activated by the addition of recombinant Prp28p and could hold valuable insight into how the spliceosome becomes active as well as definitively identifying the target of Prp28p.
212 Figure 4.1. Model for spliceosomal rearrangements leading to U1 snRNP release. A) During recognition of the 5’ SS, the U1 snRNP/5’ SS association is stabilized by the U1 snRNA/5’ SS base-pairing as well as protein factors including U1-C, Prp42p and Snu71p. At this time Prp8p may prevent Prp28p and Brr2p from recognizing their substrates. B) During spliceosome activation, a conformational change in Prp8p may allow Prp28p and Brr2p to access their substrates and may also assist in the binding of the U6 snRNA to the 5’ SS. At this time Prp28p aids in the dissociation of U1 by unwinding the RNA duplex and counteracting the stabilizing protein factors.
213 A B U1 snRNP Snu71p U1 snRNP
Snu71p Prp42p
Prp42p U1-C p Prp28p 214 U1-C p Prp28p
Prp8p U6 U6 Brr2p Brr2p Prp8p
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