Spliceosomal snRNAs are transcribed from specialized promoters, which recruit RNA polymerase II cofactors that aid in proper 3′ end maturation of these non-polyadenylated transcripts.
Like most non-coding RNAs, small nuclear RNAs (snRNAs) use cognate antisense elements to interact with their nucleic acid targets via base pairing.
Assembly of functional small nuclear ribonucleoproteins (snRNPs) involves a series of non-functional intermediates that are often sequestered in subcellular compartments that are distinct from their sites of action.
snRNP function requires multiple protein partners (such as DExD/H helicases or WD box proteins) the roles of which may include modulating RNA structure or tethering an enzyme.
snRNPs recognize specific sequences in pre-mRNAs and assemble into the spliceosome in a stepwise manner. The splicing reaction itself is catalysed by U6/U2 snRNA complex that resembles a self-splicing ribozyme.
Alternative splicing is typically regulated by multiple cis-elements and trans-factors, which form complex interaction networks that may provide a great deal of regulatory plasticity.
Pre-mRNA splicing can be regulated throughout the entire spliceosomal assembly pathway, although the early steps are the main stages of regulation.
One of the most amazing findings in molecular biology was the discovery that eukaryotic genes are discontinuous, with coding DNA being interrupted by stretches of non-coding sequence. The subsequent realization that the intervening regions are removed from pre-mRNA transcripts via the activity of a common set of small nuclear RNAs (snRNAs), which assemble together with associated proteins into a complex known as the spliceosome, was equally surprising. How do cells coordinate the assembly of this molecular machine? And how does the spliceosome accurately recognize exons and introns to carry out the splicing reaction? Insights into these questions have been gained by studying the life cycle of spliceosomal snRNAs from their transcription, nuclear export and re-import to their dynamic assembly into the spliceosome. This assembly process can also affect the regulation of alternative splicing and has implications for human disease.
Most genes in higher eukaryotes are transcribed as pre-mRNAs that contain intervening sequences (introns), as well as expressed sequences (exons). Discovered in the late 1970s, introns are now known to be removed during the process of pre-mRNA splicing, which joins exons together to produce mature mRNAs1,2. Because most human genes contain multiple introns, splicing is a crucial step in gene expression. Although the splicing reaction is chemically simple, what occurs inside a cell is much more complicated: splicing is catalysed in two distinct steps by a dynamic ribonucleoprotein (RNP) machine called the spliceosome3, requiring hydrolysis of a large quantity of ATP4. This increased complexity is thought to ensure that splicing is accurate and regulated.
The spliceosome is composed of five different RNP subunits, along with many associated protein cofactors4,5. To distinguish them from other cellular RNPs, such as ribosomal subunits, the spliceosomal subunits were termed small nuclear RNPs (snRNPs). As with ribosome assembly, the biogenesis of spliceosomal snRNPs is a multistep process that takes place in distinct subcellular compartments. A common principle in the biogenesis of snRNPs is the assembly of stable, but inactive, pre-RNPs that require maturation at locations that are distinct from their sites of function. Assembly of functional complexes and delivery to their final destinations are often regulated by progression through a series of intermediate complexes and subcellular locales.
In this Review, we discuss the key steps in the life cycle of spliceosomal snRNPs. We focus on how small nuclear RNAs (snRNAs) are synthesized and assembled with proteins into RNPs and, furthermore, how the snRNPs are assembled into the spliceosome. Finally, we highlight our current knowledge of regulatory proteins and how they affect snRNP function. We draw on recent insights from molecular, genetic, genomic and ultrastructural studies to illustrate how these factors ultimately dictate splice site choice.
Biogenesis of spliceosomal RNPs
The snRNAs are a group of abundant, non-coding, non-polyadenylated transcripts that carry out their functions in the nucleoplasm. On the basis of common sequence features and protein cofactors, they can be subdivided into two major classes: Sm and Sm-like snRNAs6. Below, we focus on the biogenesis and processing of the major and minor Sm-class spliceosomal snRNAs: U1, U2, U4, U4atac, U5, U11 and U12. Biogenesis of the Sm-like snRNAs (U6 and U6atac) is distinct from that of Sm-class RNAs6 and is not discussed in detail here.
Transcription and processing of snRNAs. In metazoans, transcription and processing of snRNAs are coupled by a cellular system that is parallel to, but distinct from, the one that generates mRNAs. Indeed, snRNA genes share many common features with protein-coding genes, including the relative positioning of elements that control transcription and RNA processing (Fig. 1). Sm-class snRNAs are transcribed from highly specialized RNA polymerase II (Pol II) promoters that contain proximal and distal sequence elements similar to the TATA box and enhancer sequences, respectively, of protein-coding genes. In addition to the general transcription factors (GTFs; consisting of transcription initiation factor IIA (TFIIA), TFIIB, TFIIE and TFIIF), initiation of snRNA transcription requires binding of a pentameric factor called the snRNA-activating protein complex (SNAPc)7,8. Promoter-swapping experiments have shown that factors required for the accurate recognition of snRNA 3′-processing signals must load onto the polymerase in a promoter-proximal manner9. Specific post-translational modifications of the carboxy-terminal domain (CTD) of the Pol II large subunit are important for loading these processing factors and for accurate processing10,11. Similar to other Pol II transcripts, capping of the 5′ end of an snRNA and cleavage of its 3′ end are thought to occur co-transcriptionally (Fig. 1).
Maturation of the snRNA 3′ end requires a large, multisubunit factor called the integrator complex12,13, which recognizes a downstream processing signal (called the 3′-box) and endonucleolytically cleaves the nascent transcript as it emerges from the polymerase (Fig. 1). Whether this cleavage occurs before, or concomitant with, the arrival of Pol II at the downstream terminator sequence is not known. Interestingly, integrator subunit 11 (INTS11) and INTS9 share important sequence similarities to components of the mRNA 3′ end-processing machinery, cleavage and polyadenylation specificity factor 73 kDa subunit (CPSF73) and CPSF100, respectively12,14,15. However, beyond these two subunits, the integrator complex proteins bear little similarity to those involved in mRNA cleavage and polyadenylation13,16. Notably, the cyclin-dependent kinase 8 (Cdk8)–cyclin C heterodimer shows snRNA 3′-processing activity in a reporter assay and physically associates with the integrator complex13. Although the kinase activity of Cdk8–cyclin C is also essential for processing, whether it phosphorylates integrator subunits and/or the Pol II CTD remains unclear13. Thus, the precise mechanism by which metazoan Pol II snRNA gene transcription is terminated remains mysterious. What is clear is that 3′-end processing of Sm-class snRNAs requires three important features: an snRNA-specific promoter, a cis-acting 3′-box element located downstream of the cleavage site and an assortment of trans-acting factors that load onto the Pol II CTD (Fig. 1).
Nuclear export, Cajal bodies and RNP quality control. Sm-class snRNPs primarily function in the nucleus. However, in most species, newly synthesized snRNAs are first exported to the cytoplasm, where they undergo additional maturation steps before they are imported back into the nucleus. Notable exceptions to this rule are found in budding yeast and trypanosomes, in which RNP assembly is thought to be entirely nuclear17,18,19,20,21. Why cells export precursor snRNAs to the cytoplasm only to re-import them after their assembly into stable RNP particles is not known. This property is not unique to snRNAs: ribosomal subunits, which function in the cytoplasm, are primarily assembled in the nucleolus22. Both types of RNP certainly undergo remodelling steps within their 'destination' compartments, but the initial stages of particle assembly take place in remote cellular locations. This arrangement provides a plausible mechanism for quality control, ensuring that partially assembled RNPs do not come into contact with their substrates.
Most types of RNA, including ribosomal RNA, tRNA, mRNA, microRNA (miRNA) and signal recognition particle (SRP) RNA, are exported to the cytoplasm following nuclear transcription and processing. Emerging evidence points to a role for nuclear RNA-binding factors in specifying the cytoplasmic fate of RNAs23. However, the connections between RNA processing and nuclear export are not as well worked out as they are for transcription and 3′-end formation. Typically, specific RNA sequences and/or structures are the determinants that promote direct or indirect binding to the appropriate transport receptor (as occurs for tRNAs and rRNAs)24. Because Sm-class snRNAs and mRNAs are both transcribed by Pol II, they share a 5′-cap structure, raising the issue of how the export machinery discriminates between these two types of RNA. Solving this long-standing riddle, an elegant series of papers has shown that snRNAs are distinguished from mRNAs on the basis of their length and their association with heterogeneous nuclear RNP (hnRNP) C1–C2 proteins25,26,27,28. Pol II mRNA transcripts that are longer than ∼250 nucleotides are bound by hnRNP C1–C2 tetramers and shunted towards the nuclear RNA export factor 1–NTF2-related export 1 (NXF1–NXT1; also known as TAP–p15) mRNA export pathway28. Transcripts shorter than this threshold are exported by a distinct set of factors that includes the cap-binding complex (cap-binding protein 80 kDa (CBP80; also known as NCBP1) and CBP20 (also known as NCBP2))29, the snRNA-specific export adaptor phosphorylated adapter RNA export (PHAX)30 and arsenite resistance 2 (ARS2; also known as SRRT)31. These proteins form a link between the 5′ cap and the export receptor chromosome region maintenance 1 (CRM1; also known as exportin 1), which interacts with nuclear pore proteins to promote export32 (Fig. 2). Although PHAX can bind to mRNA 5′ caps in vitro, it is inhibited from doing so in vivo by hnRNP C1–C2 (Ref. 28).
Several lines of evidence indicate that precursor snRNA transcripts often traffic through snRNP-rich nuclear structures known as Cajal bodies on their way out of the nucleus. First, in situ hybridization shows that pre-snRNA transcripts with long 3′ extensions localize to mammalian Cajal bodies33. Second, microinjection experiments in Xenopus laevis oocyte nuclei reveal that pre-snRNAs temporarily accumulate in Cajal bodies, and that this localization decreases over time as the RNAs are exported34. Third, PHAX and CRM1 are concentrated in Cajal bodies35,36. Fourth, inhibition of PHAX activity interferes with snRNA export30 and has been shown to cause pre-snRNAs to accumulate within frog oocyte Cajal bodies34, as well as to result in dispersal of mammalian Cajal body components37. The data are most consistent with a model in which the assembly of pre-export complexes is facilitated within Cajal bodies and is followed by nuclear export on docking to CRM1. The model further invokes a function for Cajal bodies in nuclear RNP remodelling38 and sorting23, as outlined below.
Cytoplasmic RNP assembly and the SMN complex. After the pre-snRNA translocates to the cytoplasm, dissociation of the export complex (Fig. 2) is triggered by dephosphorylation of PHAX39. The survival motor neuron (SMN) protein complex, which includes SMN and several tightly associated proteins, collectively called GEMINs40,41,42,43,44, is thought to regulate the entire cytoplasmic phase of the snRNP cycle. The SMN complex recruits the newly exported snRNAs and combines them with a set of seven Sm proteins to form a toroidal ring around an RNA-binding site that is present within each of the eponymous Sm-class snRNAs (Fig. 3). The Sm proteins are delivered to the SMN complex via the activity of the protein Arg N-methyltransferase 5 (PRMT5) complex, which methylates C-terminal arginine residues within SmB, SmD1 and SmD3 (Refs 45,46) and then chaperones delivery of partially assembled Sm subcomplexes47,48. Binding to the SMN complex is therefore proposed to initiate in the cytoplasm, and GEMIN5 is thought to be the factor responsible for recognition of Sm site-containing RNAs49. Assembly of the Sm core not only stabilizes the snRNA by protecting it from nucleases but also is required for the downstream RNA-processing steps that culminate in nuclear import. The physiological relevance of Sm core assembly has also been emphasized by the demonstration that mutations in the gene encoding the SMN protein cause the human neuromuscular disease spinal muscular atrophy (Box 1).
Sm proteins do not bind the snRNA as a pre-formed ring. Instead, they form heterodimeric (SmD1–SmD2 and SmB–SmD3) or heterotrimeric (SmE–SmF–SmG) subcomplexes (Fig. 3). When purified in vitro, these subcomplexes spontaneously coalesce into a ring only in the presence of an appropriate RNA target50,51,52. However, in cell extracts, this reaction requires the whole SMN complex as well as ATP40. In vivo, the SMN complex is thus thought to provide added specificity, to avoid assembly of Sm cores onto non-target RNAs41,49 and to accelerate formation of the final product from kinetically trapped intermediates48.
One of the most surprising insights from recent studies of the SMN complex is that the SMN protein is probably not the primary architect of Sm core RNP assembly. Two crystallographic studies demonstrated that GEMIN2, a conserved member of the SMN complex53, binds directly to five of the seven Sm proteins (Fig. 3) and holds them in the proper 'horseshoe' orientation for subsequent snRNA binding and ring closure54. These results were not predicted from earlier in vitro binding studies of GEMIN2 (Ref. 55) and were surprising because previous work on Sm binding had mainly focused on SMN itself56,57. However, given that the budding yeast genome apparently lacks SMN but contains a potential GEMIN2 orthologue55,58, the idea that GEMIN2 has a starring role in Sm core assembly is gaining considerable traction.
Precisely how SMN is involved in Sm core RNP formation is still open to debate, although RNAi analyses in metazoan cells have demonstrated that it is required59,60. Moreover, SMN–GEMIN2 heterodimers are sufficient for Sm core assembly activity in vitro53. Importantly, the assembly chaperone pICln (also known as CLNS1A) (Fig. 3) may function as an SmB–SmD3 mimic that stabilizes the pentameric Sm horseshoe structure in preparation for handover to GEMIN2 (Refs 47,48). The tudor domain of SMN contains an Sm-fold61 and is hypothesized to have a mimetic role (Fig. 3), occupying the space for SmB–SmD3 during the transition between the pICln-bound intermediate and the GEMIN2–Sm pentamer structure47. The self-oligomerization activity of SMN, contained within its C-terminal YG-box domain, is also required for Sm core formation57,60,62. It is not yet clear how the C terminus of SMN, which forms a YG-zipper motif63, interfaces with the rest of the SMN molecule and other members of the SMN complex. These and other important factors will need to be addressed by future studies.
Nuclear import and RNP remodelling. Formation of the Sm ring protects and stabilizes the snRNA and initiates downstream RNA-processing steps that culminate in nuclear import of the SMN complex (Fig. 2). As part of its overall chaperoning function, the SMN complex recruits trimethylguanosine synthase 1 (TGS1), an RNA methyltransferase that modifies the snRNA 5′ end to form a 2,2,7-trimethylguanosine (TMG) structure44. The TMG cap functions as a nuclear-localization signal64. Along with a subset of factors within the SMN complex65, the Sm core itself functions as a second, parallel nuclear-localization signal66. Concomitant with (or subsequent to) these 5′ events, the 3′ end of the snRNA is exonucleolytically trimmed to its mature length. Thus, SMN-mediated assembly of the Sm core is required for proper cytoplasmic RNP maturation in vivo.
After import back into the nucleus, TMG cap formation triggers dissociation of TGS1 from the pre-import complex (Fig. 2); this is followed by binding of Snurportin67, the snRNP-specific import adaptor, to the hypermethylated cap structure. Snurportin interacts directly with the import receptor importin-β68 to promote import, although the SMN complex (or a subcomplex thereof) is also thought to accompany newly assembled snRNPs into the nucleus65. The SMN complex does not associate with nucleus-injected (that is, 'naked') RNAs; experiments in X. laevis oocyte nuclei showed that the SMN complex interacts with microinjected snRNAs only after their export to the cytoplasm69.
When an snRNP has been imported into the nucleus, it is free to diffuse throughout the interchromatin space. SMN is thought to dissociate from the snRNP fairly soon after import, as the protein does not co-purify with mature snRNP mono-particles, spliceosomes or splicing intermediates70,71,72. In most cell types, the nuclear fraction of the SMN complex localizes primarily within Cajal bodies; however, SMN also accumulates in distinct nuclear substructures called Gemini bodies (Gems)73. Cajal bodies contain a plethora of RNAs and their associated proteins, but components of Gems have thus far been limited to consituents of the SMN complex73,74.
In mammalian cells, substantial evidence points to a role for Cajal bodies in the nucleoplasmic maturation of snRNPs following nuclear import. Newly imported Sm-class RNPs transiently accumulate in Cajal bodies before localizing in other nucleoplasmic subcompartments known as nuclear speckles (see below)75,76. In nuclear transport assays using digitonin-permeabilized cells, Snurportin 1 and partially assembled (12S) U2 snRNPs accumulate in Cajal bodies77. Additional RNP-remodelling and RNA-processing steps are thought to take place in Cajal bodies, including non-coding RNA-guided covalent modification of the snRNAs78 and binding of snRNP-specific proteins79,80. Furthermore, Cajal bodies are thought to facilitate the de novo assembly and post-splicing reassembly of U4–U6 di-snRNP and U4–U6•U5 tri-snRNP81,82,83. Given that Cajal body homeostasis is disrupted by depletion of various snRNP biogenesis factors37,60,84,85, it is perhaps surprising that snRNP trafficking through Cajal bodies is not obligatory in mice or flies86,87,88 (although it seems to be essential in fish89). Taken together, these findings strongly suggest that Cajal bodies participate in RNP biogenesis on both the outbound and inbound legs of the journey of an snRNA through the cell.
Within the nucleus, spliceosomal snRNPs and their associated cofactors (for example, SR proteins) are typically excluded from nucleoli, localizing in a punctate pattern of variably sized and irregularly shaped nuclear speckles. In fact, this speckled pattern is highly diagnostic for factors involved in pre-mRNA splicing76. Speckles are extremely dynamic nucleoplasmic domains but contain little or no DNA and are thus thought to function as storage compartments90. Most splicing activity seems to localize to the borders between speckles and the adjacent chromatin domains91,92. Precisely how snRNPs and other splicing factors are recruited from the speckles to sites of active transcription is unclear. However, when the fully assembled snRNPs are loaded onto the Pol II CTD and targeted to the site of transcription, they are then poised to carry out spliceosome assembly and pre-mRNA splicing.
Spliceosomal assembly and catalysis
Non-coding RNAs typically function as adaptors that position nucleic acid targets adjacent to an enzymatic activity that is catalysed either by the RNAs themselves or by associated proteins6. Consistent with this idea, spliceosomal snRNA function is driven by base pairing with short conserved motifs located at the junctions between the expressed exon sequences and the intervening introns of target mRNAs. The 5′ splice site (5′ss) of a pre-mRNA is present at the beginning of an intron, the 3′ss is located at the end of an intron and the branch point adenosine is usually located ∼15–50 nucleotides upstream of the 3′ss (Fig. 1b). In addition to being controlled by the primary splicing signals located at exon–intron boundaries, splice site choice is modulated by multiple cis-acting regulatory elements throughout the pre-mRNA. As outlined below, spliceosomes are assembled on their targets by a multistep process in which these cis-acting elements recruit trans-acting factors that ultimately control higher order particle assembly. For more details on splicing mechanisms, readers are referred to recent reviews4,93.
Stepwise spliceosome assembly. Although spliceosome assembly is best-understood in budding yeast, the key assembly steps are well conserved in humans. For the purposes of this Review, we refer to the names of yeast proteins. First, U1 snRNP recognizes the 5′ss via base pairing of U1 snRNA to the mRNA, forming the early complex (complex E (Fig. 4a)). In addition to recognition by base pairing, the 5′ss can be recognized by U1C, a subunit of the U1 snRNP94. This process is facilitated by the Pol II CTD, which reportedly interacts directly with U1 snRNP95,96, although the functional role of this interaction is still under debate97. The interaction between the 5′ss and U1 snRNP in complex E is ATP-independent and fairly weak; it is stabilized by other factors, such as by SR proteins98,99 and the cap-binding complex100. The 3′ss of the pre-mRNA is recognized by the U2 snRNP and associated factors, such as splicing factor 1 (SF1) and U2 auxiliary factors (U2AFs), which are also components of complex E.
In a subsequent ATP-dependent process catalysed by the DExD/H helicases pre-mRNA-processing 5 (Prp5) and Sub2, U2 snRNA recognizes sequences around the branch point adenosine and interacts with U1 snRNP to form the pre-spliceosome (complex A). Formation of an intron-spanning complex A was originally described in yeast, but more complicated assembly pathways are prevalent among higher eukaryotes. Because metazoan genes contain relatively short exons (∼50–250 nucleotides) that are separated by larger introns (up to 1,000 kb), splice sites are predominantly recognized in pairs across exons through the interaction of U1 and U2 snRNPs101,102. This process is called exon definition, and the U1–U2 snRNP complex that forms across exons is known as the exon definition complex103. In a subsequent transition step, U1 and U2 snRNPs undergo poorly understood rearrangements, forming an intron-spanning interaction known as the intron definition complex; this also brings the 5′ss, branch point and 3′ss into close proximity104. Thus, the metazoan intron definition complex is generally considered to be the equivalent of complex A in yeast, whereas the metazoan exon definition complex is similar to complex E.
Formation of the exon definition complex and the subsequent transition to the intron definition complex are intermediate stages that are crucial for regulating splicing105,106. After the assembly of complex A, the U4–U6 and U5 snRNPs are recruited as a preassembled tri-snRNP to form complex B, in a reaction catalysed by the DExD/H helicase Prp28. The resulting complex B goes through a series of compositional and conformational rearrangements to form a catalytically active complex B (complex B*). Multiple RNA helicases (Brr2, 114 kDa U5 small nuclear ribonucleoprotein component (Snu114) and Prp2) are required for the activation of complex B, resulting in rearrangements that lead to the formation of the U2–U6 snRNA structure that catalyses the splicing reaction107. The activation of complex B also unwinds the U4 and U6 snRNAs, releasing U4 and U1 from the complex108, which is thought to unmask the 5′ end of U6 snRNA.
Complex B* then completes the first catalytic step of splicing, generating complex C, which contains the free exon 1 and the intron–exon 2 lariat intermediate (Fig. 4a). Complex C undergoes additional ATP-dependent rearrangements before carrying out the second catalytic step of splicing, which is dependent on Prp8, Prp16 and synthetic lethal with U5 snRNA 7 (Slu7); this results in a post-spliceosomal complex that contains the lariat intron and spliced exons. Finally, the U2, U5 and U6 snRNPs are released from the mRNP particle and recycled for additional rounds of splicing. As with other spliceosomal rearrangement steps, release of the spliced product from the spliceosome is catalysed by the DExD/H helicase Prp22 (Refs 109,110). Disassembly of the post-catalytic spliceosome is also driven by several RNA helicases (for example, Brr2, Snu114, Prp22 and Prp43) in an ATP-dependent manner111.
Single-molecule analyses have provided additional insights into the process of spliceosome assembly. Fluorescence labelling has been used to visualize how individual spliceosomal subcomplexes sequentially associate with the pre-mRNA to generate functional spliceosomes112,113. Using purified components, these in vitro studies have shown that all of the major spliceosomal assembly steps are reversible113, including the catalytic splicing steps114. This reversibility, especially of the early steps, implies that proofreading occurs during splicing115. Commitment to splicing is thought to increase as spliceosome assembly proceeds in vitro113, consistent with the idea of a reversible stage during which partially assembled spliceosomes retain the capacity to disassemble and reassemble onto an alternative splice site. Whether splicing can be reversed in vivo is unclear, and additional studies will be required to address this point.
Aside from the traditional pathway of spliceosome assembly, at least two alternative models have been proposed. In one model, spliceosome assembly does not strictly depend on a pre-mRNA substrate, and the mRNA 5′ss can be recognized by the U1 snRNP within a penta-snRNP complex containing all five snRNPs116,117. However, this penta-snRNP observed in vitro has not been supported by studies of co-transcriptional spliceosome assembly118, and most of the evidence indicates that initial spliceosome assembly requires the presence of a 5′ss in the pre-mRNA substrate119. In the other alternative model, the U4–U6•U5 tri-snRNP can be recruited to the exon definition complex, which then can be transformed directly into a cross-intron B-like complex without prior formation of a cross-intron complex A120.
Splicing is catalysed by RNA. The spliceosome is a dynamic complex the components of which undergo multiple conformational and compositional changes during the splicing reaction. Rearrangements occur between snRNAs, spliceosomal proteins and the pre-mRNA substrate, and are required in order to generate an activated spliceosome. The snRNAs, rather than the spliceosomal proteins, are believed to provide the catalytic activity. Previous genetic and biochemical studies have established that snRNAs and substrate pre-mRNA undergo a series of dynamic base-pairing rearrangements to achieve catalysis (reviewed in Ref. 121). More recently, it was shown that the two-step splicing reaction (that is, the exchange of phosphodiester bonds) could be catalysed in a protein-free system by a U6–U2 snRNA complex that resembles a self-splicing ribozyme122,123. Indeed, structural analyses have provided information regarding atomic events within the catalytic core of the spliceosome during distinct stages of the splicing reaction124. We provide a brief overview below of how the active structure of the catalytic site is generated via RNA rearrangement (see Refs 4,93,121 for more detailed reviews).
During the early stages of spliceosomal assembly, U1 snRNA base pairs with the 5′ss. Meanwhile, U2 snRNA pairs with the branch point sequence, forming a short duplex that causes the branch-point adenosine to bulge out and present its 2′ hydroxyl group as a nucleophile (Fig. 4b). Within complex A, interactions between U1 and U2 snRNPs bring the 5′ss, the branch point and 3′ss into close proximity. Subsequently, complex A associates with the U4–U6•U5 tri-snRNP (Table 1). Recruitment of this tri-snRNP complex displaces the extensive base pairing between the U4 and U6 snRNAs and leads to the formation of new base pairs between U2 and U6107 (Fig. 4b). During this process, dissociation of U4 from U6 snRNA exposes the 5′ end of U6, which then base pairs with the 5′ss, displacing U1 snRNA (Fig. 4b).
An extensive network of base pairs is thus formed between U6 and U2 snRNA, which juxtaposes the 5′ss and the branch-point adenosine for the first catalytic step of splicing. The central region of U6 snRNA forms an intramolecular stem-loop (the U6-ISL), which is key for splicing catalysis. Recruitment of the U4–U6•U5 tri-snRNP also triggers U5 snRNA interaction with exonic sequences located near the 5′ss. This interaction is probably essential for anchoring exon 1 in proximity to the exon 2 lariat in preparation for the second catalytic step of splicing (Fig. 4). During these dynamic rearrangements, the U2–U6 complex (Fig. 4b) is thought to be the active structure that catalyses both steps of the splicing reaction. This complex shares several common structural features with the group II self-splicing introns that are found in ribozymes124,125,126, suggesting that spliceosomal catalysis might be mechanistically similar to that of ribozymes127.
In addition to base pairing among and between the snRNAs, divalent cations (for example, Mg2+) are required for pre-mRNA splicing128. These metal ions might directly participate in the catalytic reactions and/or help to maintain the active RNA conformation93. Using a 'metal rescue' strategy, U6 snRNA was shown to position the divalent metal ions to catalyse both steps of splicing by stabilizing the leaving groups127. The energy requirement for both catalytic steps of splicing is minimal, but a large amount of energy is devoted to RNA remodelling of the snRNAs. Spliceosomal remodelling is primarily catalysed by multiple DExD/H RNA helicase/ATPase129 and elongation factor G (EF-G)-like GTPase130 proteins.
Certain spliceosomal proteins may also improve the efficiency of splicing by stabilizing the RNA active site in vivo. For example, Prp8 is closely associated with the catalytic core of the spliceosome131 and is required for both its catalytic steps132. The Brr2 helicase unwinds U4–U6 snRNAs to allow U6 to pair with U2 and form the catalytically active structure. Moreover, the C-terminal tail of Prp8 can interact with Brr2 and inhibit this process133, suggesting that alternating interactions between snRNAs and proteins regulate spliceosomal activation. The second catalytic step of splicing is also thought to be promoted by proteins, including Prp16, Prp18 and Slu7. Notably, the ATP-dependent activity of Prp16 is sufficient to activate complex C for the second catalytic step of splicing134.
Most genes in higher eukaryotes undergo alternative splicing to produce multiple isoforms with distinct activities. The spliceosome is responsible for directing both constitutive and alternative splicing, and regulation of its assembly is a key control point in these processes. Alternative splicing is tightly controlled in different tissues at distinct developmental stages, and the dysregulation of splicing is associated with several human diseases (Box 1). Human introns are several to hundreds of kilobases in length (∼5 kb on average) and contain numerous 'decoy' splice sites (that is, sequences that have a similar degree of consensus matching to authentic sites). A pair of decoy splice sites often form pseudo-exons that resemble authentic exons in terms of length and splice site strength but are very rarely, if ever, spliced135. So, despite these prevalent decoy sites, the splicing process occurs with high fidelity, suggesting that additional sequence features aside from core splicing signals contribute to exon–intron definition.
Cis-acting elements regulate splicing. Alternative splicing is typically controlled by numerous cis-regulatory RNA elements that serve as either splicing enhancers or silencers. On the basis of their locations and activities, these splicing regulatory elements (SREs) are classified as exonic splicing enhancers (ESEs), intronic splicing enhancers (ISEs), exonic splicing silencers (ESSs) or intronic splicing silencers (ISSs). Although the activities of SREs are often context dependent (Fig. 5a), these sequences generally function by recruiting trans-acting splicing factors that activate or suppress different steps of the splicing reaction136,137.
How splicing factors affect splicing decisions has been a topic of extensive research. Many splicing factors are auxiliary proteins of the spliceosome and interact with its core components to regulate splicing5,138,139,140. Most known splicing factors control splicing by affecting the early and intermediate steps of spliceosomal assembly: formation of the exon definition complex and the subsequent transition to the intron-spanning complex A. A well-studied example is the protein polypyrimidine tract-binding (PTB), which typically inhibits splicing by binding to short polypyrimidine-rich elements in pre-mRNAs. When binding to exons, PTB can cause exon skipping by recognizing an ESS and inhibiting formation of the exon definition complex141. PTB can also inhibit splicing by affecting the transition from an exon definition complex to an intron definition complex106, and can directly interact with U1 snRNP to prevent its interaction with other spliceosomal components142 (Fig. 5a). Similarly, the splicing factor RNA-binding motif 5 (RBM5) interacts with a U2 snRNP component (U2AF65) and inhibits the transition from an exon definition to an intron definition complex105. In addition, hnRNP L and hnRNP A1 induce extended contacts between U1 snRNA at the 5′ss and neighbouring exonic sequences that, in turn, inhibit stable association of U6 snRNA and subsequent spliceosomal catalysis143. In addition to the early steps of spliceosomal assembly, an alternative exon in the CD45 (also known as PTPRC) mRNA was found to be inhibited after ATP-dependent exon recognition144, suggesting that alternative splicing can be regulated at many points along the spliceosomal assembly pathway.
The activities of SREs often depend on their relative locations within pre-mRNAs (Fig. 5b). This context dependence highlights the flexiblity of the interactions of the splicing regulatory factors with the core splicing machinery. Given the complexities of spliceosomes, it is not surprising that the effects of splicing factors on core spliceosomal components might vary, depending on their relative positions on the pre-mRNA. For example, oligo-G tracts commonly enhance splicing from intronic locations by recruiting hnRNP H145,146, but these same elements can inhibit splicing when located in exons147,148 (Fig. 5b). The underlying mechanism for such activities may involve inhibition of the exon definition complex by hnRNP H 'across' the site of binding. Similarly, the YCAY motifs that are recognized by the neuro-oncological ventral antigen (NOVA) family of neuron-specific splicing factors can function as ESEs, ISEs or ISSs, depending on their positions relative to the regulated exon149 (Fig. 5b). SR proteins usually promote splicing when bound to exons, but they can inhibit splicing when associated with introns150. Moreover, hnRNP A1 can inhibit splicing from either exonic or intronic locations150 (Fig. 5b). Notable exceptions to these rules have also been observed. For example, the Drosophila melanogaster orthologues of hnRNP A1 can enhance splicing from an intron151.
U1 snRNPs can also suppress splicing or inhibit polyadenylation by interacting with 5′ss-like RNA elements. In an unbiased screen, sequences resembling 5′ splice sites were identified as ESSs that inhibit exon inclusion152. The binding kinetics between U1 snRNP and the 5′ss can also affect alternative splice site choice, independently of the activities of other splicing factors153. Non-conventional functions of U1 snRNPs in preventing premature mRNA cleavage and polyadenylation are discussed in greater detail in Box 2.
Other influences on splicing. The accessibility of splice sites or cis-acting SREs can be influenced by pre-mRNA structures and binding proteins. For example, a stem-loop sequence located at the 5′ss of exon 10 of the human TAU (also known as MAPT) gene directly affects the use of the 5′ss. Stabilization of this stem-loop decreases exon 10 inclusion and, reciprocally, its destabilization increases exon 10 inclusion154. Another example is the Down syndrome cell adhesion molecule-like (Dscam) gene in D. melanogaster, in which the secondary structure of the intron ensures mutually exclusive splicing of alternative exons155,156,157. It is unclear whether examples like this are unusual cases or whether they are the general rule. Spliceosomes contain multiple DExD/H RNA helicases that can unwind RNA structures and remodel RNA–protein complexes158. Although the primary function of these helicases seems to be the rearrangement of snRNA–snRNA and snRNA–protein interactions in the spliceosome, at least one helicase (DEAD box 17 (DDX17; also known as p72)) might be able to remodel pre-mRNA structures, thus modulating alternative splicing159,160. General roles for RNA structures in splicing regulation have yet to be clearly defined, and the identification of such elements by high-throughput methods should prove very useful161.
Because splicing of most introns happens co-transcriptionally162, alternative splicing is also affected by factors that control transcription initiation and elongation. For example, the rate of transcription elongation can affect splicing events; slow elongation rates generally promote the inclusion of weak exons163,164. In addition, alternative splicing may be affected by chromatin structure and nucleosome positioning. A large number of recent reports have provided interesting insights into the connections between splicing and transcription (for further details, see Refs 165,166).
An integrated code for splicing regulation. Traditional models of splicing regulation typically consider the interaction between cis-acting SREs and their cognate factors as a one-to-one relationship. However, most splicing factors can recognize two or more SRE motifs, and each SRE motif is bound by multiple alternative factors, supporting the idea that a complex network of protein–RNA interactions is responsible for splicing regulation150,167. This pattern of overlapping binding specificities may enable a variety of regulatory relationships between splicing regulators. Multiple proteins with similar splicing regulatory activities might bind the same motif, resulting in functional redundancy; alternatively, one factor might displace another factor with opposite activity to confer functional antagonism. For example, in HeLa cells, neuronal PTB (nPTB) can compensate for depletion of PTB168, whereas during neural development replacement of PTB by nPTB is thought to initiate an alternative splicing programme169. RNA-binding factors with overlapping specificities may also provide subtle fine-tuning of splicing levels. Importantly, the densely connected network of SREs and their cognate splicing factors suggests that individual exons are often controlled by multiple factors to achieve regulatory plasticity. To assemble a set of splicing regulatory rules (known as the 'splicing code'), computational models have been applied to integrate the actions of multiple splicing factors and SREs, thereby allowing splicing outcomes to be predicted from sequence information in the pre-mRNA152,170.
Conclusions and perspectives
A major challenge in the post-genomic age of molecular biology is to understand how a limited number of human genes can generate a proteome that has five times the number of proteins171. The spliceosome, which reads the information for splicing each pre-mRNA transcript, is probably the most complicated RNA–protein complex inside the eukaryotic cell172. Although important insights have been obtained during the past decade, there are still many unanswered questions about the biogenesis of this macromolecular machine. For example, the signalling factors that regulate snRNP biogenesis are poorly understood, as are the functions of many post-translational modifications of snRNP proteins. Moreover, a key question is how conformational and compositional changes within the spliceosome dictate splicing outcomes. Detailed studies of spliceosome dynamics should provide much-needed answers.
Another important research goal is to understand the 'splicing code' by which exon inclusion or exclusion by the spliceosome is controlled in different tissues and cell types170. Recent advances in functional genomics have fuelled identification of the myriad regulatory elements and splicing factors involved, providing the research community with a near-complete 'parts list' of the splicing regulatory machinery. Integration of this information should help to determine the mechanism by which the splicing code is read by the spliceosome and ultimately provide a better understanding of complicated gene expression networks.
Berget, S. M., Moore, C. & Sharp, P. A. Spliced segments at the 5′ terminus of adenovirus 2 late mRNA. Proc. Natl Acad. Sci. USA 74, 3171–3175 (1977).
Chow, L. T., Gelinas, R. E., Broker, T. R. & Roberts, R. J. An amazing sequence arrangement at the 5′ ends of adenovirus 2 messenger RNA. Cell 12, 1–8 (1977).
Lerner, M. R., Boyle, J. A., Mount, S. M., Wolin, S. L. & Steitz, J. A. Are snRNPs involved in splicing? Nature 283, 220–224 (1980).
Will, C. L. & Luhrmann, R. Spliceosome structure and function. Cold Spring Harb. Perspect. Biol. 3, a003707 (2011).
Jurica, M. S. & Moore, M. J. Pre-mRNA splicing: awash in a sea of proteins. Mol. Cell 12, 5–14 (2003).
Matera, A. G., Terns, R. M. & Terns, M. P. Non-coding RNAs: lessons from the small nuclear and small nucleolar RNAs. Nature Rev. Mol. Cell Biol. 8, 209–220 (2007).
Henry, R. W., Mittal, V., Ma, B., Kobayashi, R. & Hernandez, N. SNAP19 mediates the assembly of a functional core promoter complex (SNAPc) shared by RNA polymerases II and III. Genes Dev. 12, 2664–2672 (1998).
Hung, K. H. & Stumph, W. E. Regulation of snRNA gene expression by the Drosophila melanogaster small nuclear RNA activating protein complex (DmSNAPc). Crit. Rev. Biochem. Mol. Biol. 46, 11–26 (2011).
Hernandez, N. & Weiner, A. M. Formation of the 3′ end of U1 snRNA requires compatible snRNA promoter elements. Cell 47, 249–258 (1986).
Egloff, S. et al. The integrator complex recognizes a new double mark on the RNA polymerase II carboxyl-terminal domain. J. Biol. Chem. 285, 20564–20569 (2010).
Egloff, S. et al. Serine-7 of the RNA polymerase II CTD is specifically required for snRNA gene expression. Science 318, 1777–1779 (2007).
Baillat, D. et al. Integrator, a multiprotein mediator of small nuclear RNA processing, associates with the C-terminal repeat of RNA polymerase II. Cell 123, 265–276 (2005). Identifies the complex that carries out pre-snRNA 3′-end processing.
Chen, J. et al. An RNAi screen identifies additional members of the Drosophila Integrator complex and a requirement for cyclin C/Cdk8 in snRNA 3′-end formation. RNA 18, 2148–2156 (2012).
Weiner, A. M. E Pluribus Unum: 3′ end formation of polyadenylated mRNAs, histone mRNAs, and U snRNAs. Mol. Cell 20, 168–170 (2005).
Mandel, C. R. et al. Polyadenylation factor CPSF-73 is the pre-mRNA 3′-end-processing endonuclease. Nature 444, 953–956 (2006).
Ezzeddine, N. et al. A subset of Drosophila integrator proteins is essential for efficient U7 snRNA and spliceosomal snRNA 3′-end formation. Mol. Cell. Biol. 31, 328–341 (2011).
Boon, K. L. et al. prp8 mutations that cause human retinitis pigmentosa lead to a U5 snRNP maturation defect in yeast. Nature Struct. Mol. Biol. 14, 1077–1083 (2007).
Murphy, M. W., Olson, B. L. & Siliciano, P. G. The yeast splicing factor Prp40p contains functional leucine-rich nuclear export signals that are essential for splicing. Genetics 166, 53–65 (2004).
Tkacz, I. D. et al. Identification of novel snRNA-specific Sm proteins that bind selectively to U2 and U4 snRNAs in Trypanosoma brucei. RNA 13, 30–43 (2007).
Palfi, Z. et al. SMN-assisted assembly of snRNP-specific Sm cores in trypanosomes. Genes Dev. 23, 1650–1664 (2009).
Jae, N. et al. snRNA-specific role of SMN in trypanosome snRNP biogenesis in vivo. RNA Biol. 8, 90–100 (2011).
Hernandez-Verdun, D., Roussel, P., Thiry, M., Sirri, V. & Lafontaine, D. L. The nucleolus: structure/function relationship in RNA metabolism. Wiley Interdiscip. Rev. RNA 1, 415–431 (2010).
Ohno, M. Size matters in RNA export. RNA Biol. 9, 1413–1417 (2012).
Cullen, B. R. Nuclear RNA export. J. Cell Sci. 116, 587–597 (2003).
Ohno, M., Segref, A., Kuersten, S. & Mattaj, I. W. Identity elements used in export of mRNAs. Mol. Cell 9, 659–671 (2002).
Masuyama, K., Taniguchi, I., Kataoka, N. & Ohno, M. RNA length defines RNA export pathway. Genes Dev. 18, 2074–2085 (2004).
Fuke, H. & Ohno, M. Role of poly (A) tail as an identity element for mRNA nuclear export. Nucleic Acids Res. 36, 1037–1049 (2008).
McCloskey, A. Taniguchi, I., Shinmyozu, K. & Ohno, M. hnRNP C tetramer measures RNA length to classify RNA polymerase II transcripts for export. Science 335, 1643–1646 (2012). First identification of a specific function for the non-shuttling hnRNP C-type proteins in RNA export.
Izaurralde, E. et al. A nuclear cap binding protein complex involved in pre-mRNA splicing. Cell 78, 657–668 (1994).
Ohno, M., Segref, A., Bachi, A., Wilm, M. & Mattaj, I. W. PHAX, a mediator of U snRNA nuclear export whose activity is regulated by phosphorylation. Cell 101, 187–198 (2000).
Hallais, M. et al. CBC–ARS2 stimulates 3′-end maturation of multiple RNA families and favors cap-proximal processing. Nature Struct. Mol. Biol. 20, 1358–1366 (2013). Shows that Ars2 forms 5′ cap-binding subcomplexes that participate in 3′-end processing of three distinct classes of transcript.
Fornerod, M., Ohno, M., Yoshida, M. & Mattaj, I. W. CRM1 is an export receptor for leucine-rich nuclear export signals. Cell 90, 1051–1060 (1997).
Smith, K. P. & Lawrence, J. B. Interactions of U2 gene loci and their nuclear transcripts with Cajal (coiled) bodies: evidence for PreU2 within Cajal bodies. Mol. Biol. Cell 11, 2987–2998 (2000).
Suzuki, T., Izumi, H. & Ohno, M. Cajal body surveillance of U snRNA export complex assembly. J. Cell Biol. 190, 603–612 (2010).
Boulon, S. et al. PHAX and CRM1 are required sequentially to transport U3 snoRNA to nucleoli. Mol. Cell 16, 777–787 (2004).
Frey, M. R. & Matera, A. G. RNA-mediated interaction of Cajal bodies and U2 snRNA genes. J. Cell Biol. 154, 499–509 (2001).
Lemm, I. et al. Ongoing U snRNP biogenesis is required for the integrity of Cajal bodies. Mol. Biol. Cell 17, 3221–3231 (2006).
Matera, A. G., Izaguire-Sierra, M., Praveen, K. & Rajendra, T. K. Nuclear bodies: random aggregates of sticky proteins or crucibles of macromolecular assembly? Dev. Cell 17, 639–647 (2009).
Kitao, S. et al. A compartmentalized phosphorylation/dephosphorylation system that regulates U snRNA export from the nucleus. Mol. Cell. Biol. 28, 487–497 (2008).
Meister, G., Buhler, D., Pillai, R., Lottspeich, F. & Fischer, U. A multiprotein complex mediates the ATP-dependent assembly of spliceosomal U snRNPs. Nature Cell Biol. 3, 945–949 (2001).
Pellizzoni, L., Yong, J. & Dreyfuss, G. Essential role for the SMN complex in the specificity of snRNP assembly. Science 298, 1775–1779 (2002).
Massenet, S., Pellizzoni, L., Paushkin, S., Mattaj, I. W. & Dreyfuss, G. The SMN complex is associated with snRNPs throughout their cytoplasmic assembly pathway. Mol. Cell. Biol. 22, 6533–6541 (2002).
Narayanan, U., Ospina, J. K., Frey, M. R., Hebert, M. D. & Matera, A. G. SMN, the spinal muscular atrophy protein, forms a pre-import snRNP complex with snurportin1 and importin β. Hum. Mol. Genet. 11, 1785–1795 (2002).
Mouaikel, J. et al. Interaction between the small-nuclear-RNA cap hypermethylase and the spinal muscular atrophy protein, survival of motor neuron. EMBO Rep. 4, 616–622 (2003).
Meister, G. et al. Methylation of Sm proteins by a complex containing PRMT5 and the putative U snRNP assembly factor pICln. Curr. Biol. 11, 1990–1994 (2001).
Friesen, W. J. et al. The methylosome, a 20S complex containing JBP1 and pICln, produces dimethylarginine-modified Sm proteins. Mol. Cell. Biol. 21, 8289–8300 (2001).
Grimm, C. et al. Structural basis of assembly chaperone-mediated snRNP formation. Mol. Cell 49, 692–703 (2013).
Chari, A. et al. An assembly chaperone collaborates with the SMN complex to generate spliceosomal snRNPs. Cell 135, 497–509 (2008).
Yong, J., Kasim, M., Bachorik, J. L., Wan, L. & Dreyfuss, G. Gemin5 delivers snRNA precursors to the SMN complex for snRNP biogenesis. Mol. Cell 38, 551–562 (2010).
Raker, V. A., Plessel, G. & Luhrmann, R. The snRNP core assembly pathway: identification of stable core protein heteromeric complexes and an snRNP subcore particle in vitro. EMBO J. 15, 2256–2269 (1996).
Kambach, C. et al. Crystal structures of two Sm protein complexes and their implications for the assembly of the spliceosomal snRNPs. Cell 96, 375–387 (1999).
Leung, A. K., Nagai, K. & Li, J. Structure of the spliceosomal U4 snRNP core domain and its implication for snRNP biogenesis. Nature 473, 536–539 (2011). Co-crystal structure of U4 snRNA construct with an Sm core definitively shows that the RNA passes through the hole in the Sm ring.
Kroiss, M. et al. Evolution of an RNP assembly system: a minimal SMN complex facilitates formation of UsnRNPs in Drosophila melanogaster. Proc. Natl Acad. Sci. USA 105, 10045–10050 (2008). Shows that both human and fruitfly SMN–GEMIN2 heterodimers are sufficient for mediating Sm core assembly in vitro.
Zhang, R. et al. Structure of a key intermediate of the SMN complex reveals Gemin2's crucial function in snRNP assembly. Cell 146, 384–395 (2011). Together with reference 46, these papers identify key intermediates in the Sm core assembly pathway, highlighting an unexpected role for GEMIN2.
Liu, Q., Fischer, U., Wang, F. & Dreyfuss, G. The spinal muscular atrophy disease gene product, SMN, and its associated protein SIP1 are in a complex with spliceosomal snRNP proteins. Cell 90, 1013–1021 (1997).
Buhler, D., Raker, V., Luhrmann, R. & Fischer, U. Essential role for the tudor domain of SMN in spliceosomal U snRNP assembly: implications for spinal muscular atrophy. Hum. Mol. Genet. 8, 2351–2357 (1999).
Pellizzoni, L., Charroux, B. & Dreyfuss, G. SMN mutants of spinal muscular atrophy patients are defective in binding to snRNP proteins. Proc. Natl Acad. Sci. USA 96, 11167–11172 (1999).
Hannus, S., Buhler, D., Romano, M., Seraphin, B. & Fischer, U. The Schizosaccharomyces pombe protein Yab8p and a novel factor, Yip1p, share structural and functional similarity with the spinal muscular atrophy-associated proteins SMN and SIP1. Hum. Mol. Genet. 9, 663–674 (2000).
Rajendra, T. K. et al. A Drosophila melanogaster model of spinal muscular atrophy reveals a function for SMN in striated muscle. J. Cell Biol. 176, 831–841 (2007).
Shpargel, K. B. & Matera, A. G. Gemin proteins are required for efficient assembly of Sm-class ribonucleoproteins. Proc. Natl Acad. Sci. USA 102, 17372–17377 (2005). Assays individual Gemins, as well as a panel of SMN missense mutants for ability to carry out Sm core assembly, showing that certain SMA-causing alleles are functional, whereas others are not.
Selenko, P. et al. SMN Tudor domain structure and its interaction with the Sm proteins. Nature Struct. Biol. 8, 27–31 (2001).
Lorson, C. L. et al. SMN oligomerization defect correlates with spinal muscular atrophy severity. Nature Genet. 19, 63–66 (1998).
Martin, R., Gupta, K., Ninan, N. S., Perry, K. & Van Duyne, G. D. The survival motor neuron protein forms soluble glycine zipper oligomers. Structure 20, 1929–1939 (2012).
Fischer, U. & Luhrmann, R. An essential signaling role for the m3G cap in the transport of U1 snRNP to the nucleus. Science 249, 786–790 (1990).
Narayanan, U., Achsel, T., Luhrmann, R. & Matera, A. G. Coupled in vitro import of U snRNPs and SMN, the spinal muscular atrophy protein. Mol. Cell 16, 223–234 (2004).
Fischer, U., Sumpter, V., Sekine, M., Satoh, T. & Luhrmann, R. Nucleo-cytoplasmic transport of U snRNPs: definition of a nuclear location signal in the Sm core domain that binds a transport receptor independently of the m3G cap. EMBO J. 12, 573–583 (1993).
Huber, J. et al. Snurportin1, an m3G-cap-specific nuclear import receptor with a novel domain structure. EMBO J. 17, 4114–4126 (1998).
Palacios, I., Hetzer, M., Adam, S. A. & Mattaj, I. W. Nuclear import of U snRNPs requires importin β. EMBO J. 16, 6783–6792 (1997).
Fischer, U., Liu, Q. & Dreyfuss, G. The SMN–SIP1 complex has an essential role in spliceosomal snRNP biogenesis. Cell 90, 1023–1029 (1997).
Neubauer, G. et al. Mass spectrometry and EST-database searching allows characterization of the multi-protein spliceosome complex. Nature Genet. 20, 46–50 (1998).
Trinkle-Mulcahy, L. et al. Identifying specific protein interaction partners using quantitative mass spectrometry and bead proteomes. J. Cell Biol. 183, 223–239 (2008).
Herold, N. et al. Conservation of the protein composition and electron microscopy structure of Drosophila melanogaster and human spliceosomal complexes. Mol. Cell. Biol. 29, 281–301 (2009).
Matera, A. G. & Shpargel, K. B. Pumping RNA: nuclear bodybuilding along the RNP pipeline. Curr. Opin. Cell Biol. 18, 317–324 (2006).
Stanek, D. & Neugebauer, K. M. The Cajal body: a meeting place for spliceosomal snRNPs in the nuclear maze. Chromosoma 115, 343–354 (2006).
Sleeman, J. E. & Lamond, A. I. Newly assembled snRNPs associate with coiled bodies before speckles, suggesting a nuclear snRNP maturation pathway. Curr. Biol. 9, 1065–1074 (1999).
Lamond, A. I. & Spector, D. L. Nuclear speckles: a model for nuclear organelles. Nature Rev. Mol. Cell Biol. 4, 605–612 (2003).
Ospina, J. K. et al. Cross-talk between snurportin1 subdomains. Mol. Biol. Cell 16, 4660–4671 (2005).
Jady, B. E. et al. Modification of Sm small nuclear RNAs occurs in the nucleoplasmic Cajal body following import from the cytoplasm. EMBO J. 22, 1878–1888 (2003).
Nesic, D., Tanackovic, G. & Kramer, A. A role for Cajal bodies in the final steps of U2 snRNP biogenesis. J. Cell Sci. 117, 4423–4433 (2004).
Schaffert, N., Hossbach, M., Heintzmann, R., Achsel, T. & Luhrmann, R. RNAi knockdown of hPrp31 leads to an accumulation of U4/U6 di-snRNPs in Cajal bodies. EMBO J. 23, 3000–3009 (2004).
Novotny, I., Blazikova, M., Stanek, D., Herman, P. & Malinsky, J. In vivo kinetics of U4/U6. U5 tri-snRNP formation in Cajal bodies. Mol. Biol. Cell 22, 513–523 (2011).
Stanek, D. & Neugebauer, K. M. Detection of snRNP assembly intermediates in Cajal bodies by fluorescence resonance energy transfer. J. Cell Biol. 166, 1015–1025 (2004).
Stanek, D., Rader, S. D., Klingauf, M. & Neugebauer, K. M. Targeting of U4/U6 small nuclear RNP assembly factor SART3/p110 to Cajal bodies. J. Cell Biol. 160, 505–516 (2003).
Strzelecka, M., Oates, A. C. & Neugebauer, K. M. Dynamic control of Cajal body number during zebrafish embryogenesis. Nucleus 1, 96–108 (2010).
Takata, H., Nishijima, H., Maeshima, K. & Shibahara, K. The integrator complex is required for integrity of Cajal bodies. J. Cell Sci. 125, 166–175 (2012).
Tucker, K. E. et al. Residual Cajal bodies in coilin knockout mice fail to recruit Sm snRNPs and SMN, the spinal muscular atrophy gene product. J. Cell Biol. 154, 293–307 (2001).
Liu, J. L. et al. Coilin is essential for Cajal body organization in Drosophila melanogaster. Mol. Biol. Cell 20, 1661–1670 (2009).
Walker, M. P., Tian, L. & Matera, A. G. Reduced viability, fertility and fecundity in mice lacking the cajal body marker protein, coilin. PLoS ONE 4, e6171 (2009).
Strzelecka, M. et al. Coilin-dependent snRNP assembly is essential for zebrafish embryogenesis. Nature Struct. Mol. Biol. 17, 403–409 (2010).
Spector, D. L. & Lamond, A. I. Nuclear speckles. Cold Spring Harb. Perspect. Biol. 3, a000646 (2011).
Hall, L. L., Smith, K. P., Byron, M. & Lawrence, J. B. Molecular anatomy of a speckle. Anat. Rec. A Discov. Mol. Cell. Evol. Biol. 288, 664–675 (2006).
Girard, C. et al. Post-transcriptional spliceosomes are retained in nuclear speckles until splicing completion. Nature Commun. 3, 994 (2012).
Valadkhan, S. Role of the snRNAs in spliceosomal active site. RNA Biol. 7, 345–353 (2010).
Du, H. & Rosbash, M. The U1 snRNP protein U1C recognizes the 5′ splice site in the absence of base pairing. Nature 419, 86–90 (2002).
Wiesner, S., Stier, G., Sattler, M. & Macias, M. J. Solution structure and ligand recognition of the WW domain pair of the yeast splicing factor Prp40. J. Mol. Biol. 324, 807–822 (2002).
Morris, D. P. & Greenleaf, A. L. The splicing factor, Prp40, binds the phosphorylated carboxyl-terminal domain of RNA polymerase II. J. Biol. Chem. 275, 39935–39943 (2000).
Gornemann, J. et al. Cotranscriptional spliceosome assembly and splicing are independent of the Prp40p WW domain. RNA 17, 2119–2129 (2011).
Staknis, D. & Reed, R. SR proteins promote the first specific recognition of Pre-mRNA and are present together with the U1 small nuclear ribonucleoprotein particle in a general splicing enhancer complex. Mol. Cell. Biol. 14, 7670–7682 (1994).
Cho, S. et al. Interaction between the RNA binding domains of Ser-Arg splicing factor 1 and U1–70K snRNP protein determines early spliceosome assembly. Proc. Natl Acad. Sci. USA 108, 8233–8238 (2011).
Pabis, M. et al. The nuclear cap-binding complex interacts with the U4/U6. U5 tri-snRNP and promotes spliceosome assembly in mammalian cells. RNA 19, 1054–1063 (2013).
Fox-Walsh, K. L. et al. The architecture of pre-mRNAs affects mechanisms of splice-site pairing. Proc. Natl Acad. Sci. USA 102, 16176–16181 (2005).
Xiao, X., Wang, Z., Jang, M. & Burge, C. B. Coevolutionary networks of splicing cis-regulatory elements. Proc. Natl Acad. Sci. USA 104, 18583–18588 (2007).
Sterner, D. A., Carlo, T. & Berget, S. M. Architectural limits on split genes. Proc. Natl Acad. Sci. USA 93, 15081–15085 (1996).
De Conti, L., Baralle, M. & Buratti, E. Exon and intron definition in pre-mRNA splicing. Wiley Interdiscip. Rev. RNA 4, 49–60 (2013).
Bonnal, S. et al. RBM5/Luca-15/H37 regulates Fas alternative splice site pairing after exon definition. Mol. Cell 32, 81–95 (2008).
Sharma, S., Kohlstaedt, L. A., Damianov, A., Rio, D. C. & Black, D. L. Polypyrimidine tract binding protein controls the transition from exon definition to an intron defined spliceosome. Nature Struct. Mol. Biol. 15, 183–191 (2008). Demonstrates that an early step in spliceosome assembly (transition from exon definition to intron definition complex) is a key stage for splicing regulation.
Sun, J. S. & Manley, J. L. A novel U2–U6 snRNA structure is necessary for mammalian mRNA splicing. Genes Dev. 9, 843–854 (1995).
Raghunathan, P. L. & Guthrie, C. RNA unwinding in U4/U6 snRNPs requires ATP hydrolysis and the DEIH-box splicing factor Brr2. Curr. Biol. 8, 847–855 (1998).
Ilagan, J. O., Chalkley, R. J., Burlingame, A. L. & Jurica, M. S. Rearrangements within human spliceosomes captured after exon ligation. RNA 19, 400–412 (2013)
Schwer, B. & Gross, C. H. Prp22, a DExH-box RNA helicase, plays two distinct roles in yeast pre-mRNA splicing. EMBO J. 17, 2086–2094 (1998).
Fourmann, J. B. et al. Dissection of the factor requirements for spliceosome disassembly and the elucidation of its dissociation products using a purified splicing system. Genes Dev. 27, 413–428 (2013).
Abelson, J. et al. Conformational dynamics of single pre-mRNA molecules during in vitro splicing. Nature Struct. Mol. Biol. 17, 504–512 (2010).
Hoskins, A. A. et al. Ordered and dynamic assembly of single spliceosomes. Science 331, 1289–1295 (2011).
Tseng, C. K. & Cheng, S. C. Both catalytic steps of nuclear pre-mRNA splicing are reversible. Science 320, 1782–1784 (2008).
Yang, F. et al. Splicing proofreading at 5′ splice sites by ATPase Prp28p. Nucleic Acids Res. 41, 4660–4670 (2013).
Malca, H., Shomron, N. & Ast, G. The U1 snRNP base pairs with the 5′ splice site within a penta–snRNP complex. Mol. Cell. Biol. 23, 3442–3455 (2003).
Stevens, S. W. et al. Composition and functional characterization of the yeast spliceosomal penta–snRNP. Mol. Cell 9, 31–44 (2002).
Gornemann, J., Kotovic, K. M., Hujer, K. & Neugebauer, K. M. Cotranscriptional spliceosome assembly occurs in a stepwise fashion and requires the cap binding complex. Mol. Cell 19, 53–63 (2005). Development of a novel chromatin immunoprecipitation assay to investigate co-transcriptional spliceosome assembly, demonstrating a role for the CBC in recruitment of snRNPs to nascent pre-mRNA transcripts.
Behzadnia, N., Hartmuth, K., Will, C. L. & Luhrmann, R. Functional spliceosomal A complexes can be assembled in vitro in the absence of a penta–snRNP. RNA 12, 1738–1746 (2006).
Schneider, M. et al. Exon definition complexes contain the tri-snRNP and can be directly converted into B-like precatalytic splicing complexes. Mol. Cell 38, 223–235 (2010). Together with reference 116, these studies suggest the existence of alternative spliceosome assembly pathways.
Madhani, H. D. & Guthrie, C. Dynamic RNA–RNA interactions in the spliceosome. Annu. Rev. Genet. 28, 1–26 (1994).
Valadkhan, S., Mohammadi, A., Wachtel, C. & Manley, J. L. Protein-free spliceosomal snRNAs catalyze a reaction that resembles the first step of splicing. RNA 13, 2300–2311 (2007).
Valadkhan, S., Mohammadi, A., Jaladat, Y. & Geisler, S. Protein-free small nuclear RNAs catalyze a two-step splicing reaction. Proc. Natl Acad. Sci. USA 106, 11901–11906 (2009). Together with reference 122, demonstrates that protein-free U6/U2 snRNA constructs can recognize 5′ splice site and branch point sequence to carry out the first and second steps of splicing.
Marcia, M. & Pyle, A. M. Visualizing group II intron catalysis through the stages of splicing. Cell 151, 497–507 (2012).
Toor, N., Keating, K. S. & Pyle, A. M. Structural insights into RNA splicing. Curr. Opin. Struct. Biol. 19, 260–266 (2009).
Toor, N., Keating, K. S., Taylor, S. D. & Pyle, A. M. Crystal structure of a self-spliced group II intron. Science 320, 77–82 (2008).
Fica, S. M. et al. RNA catalyses nuclear pre-mRNA splicing. Nature 503, 229–234 (2013).
Butcher, S. E. The spliceosome and its metal ions. Met. Ions Life Sci. 9, 235–251 (2011).
Cordin, O., Hahn, D. & Beggs, J. D. Structure, function and regulation of spliceosomal RNA helicases. Curr. Opin. Cell Biol. 24, 431–438 (2012).
Small, E. C., Leggett, S. R., Winans, A. A. & Staley, J. P. The EF-G-like GTPase Snu114p regulates spliceosome dynamics mediated by Brr2p, a DExD/H box ATPase. Mol. Cell 23, 389–399 (2006).
Galej, W. P., Oubridge, C., Newman, A. J. & Nagai, K. Crystal structure of Prp8 reveals active site cavity of the spliceosome. Nature 493, 638–643 (2013).
Schellenberg, M. J. et al. A conformational switch in PRP8 mediates metal ion coordination that promotes pre-mRNA exon ligation. Nature Struct. Mol. Biol. 20, 728–734 (2013).
Mozaffari-Jovin, S. et al. Inhibition of RNA helicase Brr2 by the C-terminal tail of the spliceosomal protein Prp8. Science 341, 80–84 (2013).
Ohrt, T. et al. Molecular dissection of step 2 catalysis of yeast pre-mRNA splicing investigated in a purified system. RNA 19, 902–915 (2013).
Sun, H. & Chasin, L. A. Multiple splicing defects in an intronic false exon. Mol. Cell. Biol. 20, 6414–6425 (2000).
Matlin, A. J., Clark, F. & Smith, C. W. Understanding alternative splicing: towards a cellular code. Nature Rev. Mol. Cell Biol. 6, 386–398 (2005).
Wang, Z. & Burge, C. B. Splicing regulation: from a parts list of regulatory elements to an integrated splicing code. RNA 14, 802–813 (2008).
Bessonov, S., Anokhina, M., Will, C. L., Urlaub, H. & Luhrmann, R. Isolation of an active step I spliceosome and composition of its RNP core. Nature 452, 846–850 (2008).
Zhou, Z., Licklider, L. J., Gygi, S. P. & Reed, R. Comprehensive proteomic analysis of the human spliceosome. Nature 419, 182–185 (2002). Identifies more than 100 proteins in the active spliceosome, many more than the known protein components of snRNPs.
Hegele, A. et al. Dynamic protein–protein interaction wiring of the human spliceosome. Mol. Cell 45, 567–580 (2012).
Izquierdo, J. M. et al. Regulation of Fas alternative splicing by antagonistic effects of TIA-1 and PTB on exon definition. Mol. Cell 19, 475–484 (2005).
Sharma, S., Maris, C., Allain, F. H. & Black, D. L. U1 snRNA directly interacts with polypyrimidine tract-binding protein during splicing repression. Mol. Cell 41, 579–588 (2011).
Chiou, N. T., Shankarling, G. & Lynch, K. W. HnRNP L and hnRNP A1 induce extended U1 snRNA interactions with an exon to repress spliceosome assembly. Mol. Cell 49, 972–982 (2013).
House, A. E. & Lynch, K. W. An exonic splicing silencer represses spliceosome assembly after ATP-dependent exon recognition. Nature Struct. Mol. Biol. 13, 937–944 (2006).
McCullough, A. J. & Berget, S. M. G triplets located throughout a class of small vertebrate introns enforce intron borders and regulate splice site selection. Mol. Cell. Biol. 17, 4562–4571 (1997).
Chou, M. Y., Rooke, N., Turck, C. W. & Black, D. L. hnRNP H is a component of a splicing enhancer complex that activates a c-src alternative exon in neuronal cells. Mol. Cell. Biol. 19, 69–77 (1999).
Chen, C. D., Kobayashi, R. & Helfman, D. M. Binding of hnRNP H to an exonic splicing silencer is involved in the regulation of alternative splicing of the rat β-tropomyosin gene. Genes Dev. 13, 593–606 (1999).
Caputi, M. & Zahler, A. M. Determination of the RNA binding specificity of the heterogeneous nuclear ribonucleoprotein (hnRNP) H/H′/F/2H9 family. J. Biol. Chem. 276, 43850–43859 (2001).
Ule, J. et al. An RNA map predicting Nova-dependent splicing regulation. Nature 444, 580–586 (2006).
Wang, Y. et al. A complex network of factors with overlapping affinities represses splicing through intronic elements. Nature Struct. Mol. Biol. 20, 36–45 (2013). Suggests that interactions between various cis -acting elements and trans -acting factors form a complex network that controls context-dependent splicing.
Borah, S., Wong, A. C. & Steitz, J. A. Drosophila hnRNP A1 homologs Hrp36/Hrp38 enhance U2-type versus U12-type splicing to regulate alternative splicing of the prospero twintron. Proc. Natl Acad. Sci. USA 106, 2577–2582 (2009).
Wang, Z. et al. Systematic identification and analysis of exonic splicing silencers. Cell 119, 831–845 (2004).
Yu, Y. et al. Dynamic regulation of alternative splicing by silencers that modulate 5′ splice site competition. Cell 135, 1224–1236 (2008).
Donahue, C. P., Muratore, C., Wu, J. Y., Kosik, K. S. & Wolfe, M. S. Stabilization of the tau exon 10 stem loop alters pre-mRNA splicing. J. Biol. Chem. 281, 23302–23306 (2006).
Graveley, B. R. Mutually exclusive splicing of the insect Dscam pre-mRNA directed by competing intronic RNA secondary structures. Cell 123, 65–73 (2005). A great example of how RNA structures can have a leading role in controlling a complicated regimen of mutally exclusive splicing.
Yang, Y. et al. RNA secondary structure in mutually exclusive splicing. Nature Struct. Mol. Biol. 18, 159–168 (2011).
Wang, X. et al. An RNA architectural locus control region involved in Dscam mutually exclusive splicing. Nature Commun. 3, 1255 (2012).
Bleichert, F. & Baserga, S. J. The long unwinding road of RNA helicases. Mol. Cell 27, 339–352 (2007).
Honig, A., Auboeuf, D., Parker, M. M., O'Malley, B. W. & Berget, S. M. Regulation of alternative splicing by the ATP-dependent DEAD-box RNA helicase p72. Mol. Cell. Biol. 22, 5698–5707 (2002).
Lee, C. G. RH70, a bidirectional RNA helicase, co-purifies with U1snRNP. J. Biol. Chem. 277, 39679–39683 (2002).
Weeks, K. M. Advances in RNA structure analysis by chemical probing. Curr. Opin. Struct. Biol. 20, 295–304 (2010).
Khodor, Y. L. et al. Nascent-seq indicates widespread cotranscriptional pre-mRNA splicing in Drosophila. Genes Dev. 25, 2502–2512 (2011).
Ip, J. Y. et al. Global impact of RNA polymerase II elongation inhibition on alternative splicing regulation. Genome Res. 21, 390–401 (2011).
Roberts, G. C., Gooding, C., Mak, H. Y., Proudfoot, N. J. & Smith, C. W. Co-transcriptional commitment to alternative splice site selection. Nucleic Acids Res. 26, 5568–5572 (1998).
Kornblihtt, A. R. et al. Alternative splicing: a pivotal step between eukaryotic transcription and translation. Nature Rev. Mol. Cell Biol. 14, 153–165 (2013).
Brugiolo, M., Herzel, L. & Neugebauer, K. M. Counting on co-transcriptional splicing. F1000Prime Rep. 5, 9 (2013).
Wang, Y., Ma, M., Xiao, X. & Wang, Z. Intronic splicing enhancers, cognate splicing factors and context-dependent regulation rules. Nature Struct. Mol. Biol. 19, 1044–1052 (2012).
Spellman, R., Llorian, M. & Smith, C. W. Crossregulation and functional redundancy between the splicing regulator PTB and its paralogs nPTB and ROD1. Mol. Cell 27, 420–434 (2007).
Boutz, P. L. et al. A post-transcriptional regulatory switch in polypyrimidine tract-binding proteins reprograms alternative splicing in developing neurons. Genes Dev. 21, 1636–1652 (2007).
Barash, Y. et al. Deciphering the splicing code. Nature 465, 53–59 (2010).
Nilsen, T. W. & Graveley, B. R. Expansion of the eukaryotic proteome by alternative splicing. Nature 463, 457–463 (2010).
Nilsen, T. W. The spliceosome: the most complex macromolecular machine in the cell? BioEssays 25, 1147–1149 (2003).
Singh, R. K. & Cooper, T. A. Pre-mRNA splicing in disease and therapeutics. Trends Mol. Med. 18, 472–482 (2012).
Padgett, R. A. New connections between splicing and human disease. Trends Genet. 28, 147–154 (2012).
Tanackovic, G. et al. PRPF mutations are associated with generalized defects in spliceosome formation and pre-mRNA splicing in patients with retinitis pigmentosa. Hum. Mol. Genet. 20, 2116–2130 (2011).
Utz, V. M., Beight, C. D., Marino, M. J., Hagstrom, S. A. & Traboulsi, E. I. Autosomal dominant retinitis pigmentosa secondary to pre-mRNA splicing-factor gene PRPF31 (RP11): review of disease mechanism and report of a family with a novel 3-base pair insertion. Ophthalm. Genet. 34, 183–188 (2013).
Pena, V., Liu, S., Bujnicki, J. M., Luhrmann, R. & Wahl, M. C. Structure of a multipartite protein–protein interaction domain in splicing factor prp8 and its link to retinitis pigmentosa. Mol. Cell 25, 615–624 (2007).
He, H. et al. Mutations in U4atac snRNA, a component of the minor spliceosome, in the developmental disorder MOPD I. Science 332, 238–240 (2011).
Lorson, C. L., Hahnen, E., Androphy, E. J. & Wirth, B. A single nucleotide in the SMN gene regulates splicing and is responsible for spinal muscular atrophy. Proc. Natl Acad. Sci. USA 96, 6307–6311 (1999).
Schrank, B. et al. Inactivation of the survival motor neuron gene, a candidate gene for human spinal muscular atrophy, leads to massive cell death in early mouse embryos. Proc. Natl Acad. Sci. USA 94, 9920–9925 (1997).
Gabanella, F. et al. Ribonucleoprotein assembly defects correlate with spinal muscular atrophy severity and preferentially affect a subset of spliceosomal snRNPs. PLoS ONE 2, e921 (2007).
Praveen, K., Wen, Y. & Matera, A. G. A. Drosophila model of spinal muscular atrophy uncouples snRNP biogenesis functions of survival motor neuron from locomotion and viability defects. Cell Rep. 1, 624–631 (2012).
Garcia, E. L., Lu, Z., Meers, M. P., Praveen, K. & Matera, A. G. Developmental arrest of Drosophila survival motor neuron (Smn) mutants accounts for differences in expression of minor intron-containing genes. RNA 19, 1510–1516 (2013).
Baumer, D. et al. Alternative splicing events are a late feature of pathology in a mouse model of spinal muscular atrophy. PLoS Genet. 5, e1000773 (2009). Together with references 182 and 183, these studies show that SMA phenotypes can be uncoupled from global splicing deficits. Using a missense allele that is active in Sm core assembly, reference 184 reveals a separation of SMN functions.
Cazzola, M., Rossi, M. & Malcovati, L. Biologic and clinical significance of somatic mutations of SF3B1 in myeloid and lymphoid neoplasms. Blood 121, 260–269 (2013).
Yoshida, K. et al. Frequent pathway mutations of splicing machinery in myelodysplasia. Nature 478, 64–69 (2011).
Chesnais, V. et al. Spliceosome mutations in myelodysplastic syndromes and chronic myelomonocytic leukemia. Oncotarget 3, 1284–1293 (2012).
Dhir, A., Buratti, E., van Santen, M. A., Luhrmann, R. & Baralle, F. E. The intronic splicing code: multiple factors involved in ATM pseudoexon definition. EMBO J. 29, 749–760 (2010).
Lewandowska, M. A., Stuani, C., Parvizpur, A., Baralle, F. E. & Pagani, F. Functional studies on the ATM intronic splicing processing element. Nucleic Acids Res. 33, 4007–4015 (2005).
Pagani, F. et al. A new type of mutation causes a splicing defect in ATM. Nature Genet. 30, 426–429 (2002).
Gunderson, S. I., Polycarpou-Schwarz, M. & Mattaj, I. W. U1 snRNP inhibits pre-mRNA polyadenylation through a direct interaction between U1 70K and poly(A) polymerase. Mol. Cell 1, 255–264 (1998).
Langemeier, J., Radtke, M. & Bohne, J. U1 snRNP-mediated poly(A) site suppression: beneficial and deleterious for mRNA fate. RNA Biol. 10, 180–184 (2013).
Kaida, D. et al. U1 snRNP protects pre-mRNAs from premature cleavage and polyadenylation. Nature 468, 664–668 (2010).
Almada, A. E., Wu, X., Kriz, A. J., Burge, C. B. & Sharp, P. A. Promoter directionality is controlled by U1 snRNP and polyadenylation signals. Nature 499, 360–363 (2013).
Berg, M. G. et al. U1 snRNP determines mRNA length and regulates isoform expression. Cell 150, 53–64 (2012). Together with reference 193, these genome-wide analyses illustrate a pervasive, non-splicing role for U1 snRNP in selection of the site of pre-mRNA 3′-end cleavage and polyadenylation.
Peterson, M. L., Bingham, G. L. & Cowan, C. Multiple features contribute to the use of the immunoglobulin M secretion-specific poly(A) signal but are not required for developmental regulation. Mol. Cell. Biol. 26, 6762–6771 (2006).
Hall-Pogar, T., Liang, S., Hague, L. K. & Lutz, C. S. Specific trans-acting proteins interact with auxiliary RNA polyadenylation elements in the COX-2 3′-UTR. RNA 13, 1103–1115 (2007).
Luo, W. et al. The conserved intronic cleavage and polyadenylation site of CstF-77 gene imparts control of 3′ end processing activity through feedback autoregulation and by U1 snRNP. PLoS Genet. 9, e1003613 (2013).
Michaeli, S. Trans-splicing in trypanosomes: machinery and its impact on the parasite transcriptome. Future Microbiol. 6, 459–474 (2011).
Lasda, E. L. & Blumenthal, T. Trans-splicing. Wiley Interdiscip. Rev. RNA 2, 417–434 (2011).
Bruzik, J. P. & Maniatis, T. Spliced leader RNAs from lower eukaryotes are trans-spliced in mammalian cells. Nature 360, 692–695 (1992).
Smith, E. R. et al. The little elongation complex regulates small nuclear RNA transcription. Mol. Cell 44, 954–965 (2011).
Fabrizio, P. et al. The evolutionarily conserved core design of the catalytic activation step of the yeast spliceosome. Mol. Cell 36, 593–608 (2009).
Research in the authors' laboratories is supported by US National Institutes of Health grants R01-GM053034 and R01-NS041617 (to A.G.M.), as well as R01-CA158283 and R21-AR061640 (to Z.W.). The authors apologize to those whose work could not be discussed owing to space limitations.
The authors declare no competing financial interests.
- Splice site
The short sequences at exon–intron junctions of pre-mRNA, which include the 5′ splice (splice donor) site and the 3′ splice (splice acceptor) site located at the beginning and the end of an intron, respectively.
- Heterogeneous nuclear RNP
(hnRNP). A diverse class of ribonucleoproteins (RNPs) located in the cell nucleus, and primarily involved in post-transcriptional regulation of mRNAs. The hnRNP proteins are a class of RNA-binding factors, many of which shuttle between the nucleus and cytoplasm, that are involved in regulating the processing, stability and subcellular transport of mRNPs.
- Cajal bodies
Nuclear substructures that are highly enriched in pre-mRNA splicing factors. They are thought to function as sites of ribonucleoprotein assembly and remodelling.
- Tudor domain
A conserved protein structural motif that is thought to bind to methylated arginine or lysine residues, promoting physical interactions with its target protein.
- Nuclear speckles
Sub-nuclear structures highly enriched in pre-mRNA-splicing factors. At the ultrastructural level, they correspond to domains known as interchromatin granule clusters.
- SR proteins
Proteins that contain a domain with repeats of serine (S) and arginine (R) residues and one or more RNA-recognition motifs. SR proteins are best known for their ability to bind exonic splicing enhancers and activate splicing, although some SR proteins also regulate transcription.
- Branch point
A loosely conserved short sequence usually located ∼15–50 nucleotides upstream of the 3′ splice site, before a region rich in pyrimidines (cytosine and uracil). Most branch points include an adenine nucleotide as the site of lariat formation.
- Exon definition
One of two different modes of initial splice site pairing at the early stage of splicing (the other being intron definition). During exon definition, the U1 and U2 small nuclear ribonucleoproteins (snRNPs) interact to pair the splice sites across an exon. For some small introns, the U1 and U2 snRNPs interact to pair the splice sites across introns.
About this article
Cite this article
Matera, A., Wang, Z. A day in the life of the spliceosome. Nat Rev Mol Cell Biol 15, 108–121 (2014). https://doi.org/10.1038/nrm3742
Nature Communications (2020)
SPLICELECT™: an adaptable cell surface display technology based on alternative splicing allowing the qualitative and quantitative prediction of secreted product at a single-cell level
Cellular and Molecular Neurobiology (2020)
Exonic rearrangements in DMD in Chinese Han individuals affected with Duchenne and Becker muscular dystrophies
Human Mutation (2019)