Rearrangements within human spliceosomes captured after exon ligation.

In spliceosomes, dynamic RNA/RNA and RNA/protein interactions position the pre-mRNA substrate for the two chemical steps of splicing. Not all of these interactions have been characterized, in part because it has not been possible to arrest the complex at clearly defined states relative to chemistry. Previously, it was shown in yeast that the DEAD/H-box protein Prp22 requires an extended 3' exon to promote mRNA release from the spliceosome following second-step chemistry. In line with that observation, we find that shortening the 3' exon blocks cleaved lariat intron and mRNA release in human splicing extracts, which allowed us to stall human spliceosomes in a new post-catalytic complex (P complex). In comparison to C complex, which is blocked at a point following first-step chemistry, we detect specific differences in RNA substrate interactions near the splice sites. These differences include extended protection across the exon junction and changes in protein crosslinks to specific sites in the 5' and 3' exons. Using selective reaction monitoring (SRM) mass spectrometry, we quantitatively compared P and C complex proteins and observed enrichment of SF3b components and loss of the putative RNA-dependent ATPase DHX35. Electron microscopy revealed similar structural features for both complexes. Notably, additional density is present when complexes are chemically fixed, which reconciles our results with previously reported C complex structures. Our ability to compare human spliceosomes before and after second-step chemistry has opened a new window to rearrangements near the active site of spliceosomes, which may play roles in exon ligation and mRNA release

Sample processing obscures cancer-specific alterations in leukemic transcriptomes

Substantial effort is currently devoted to identifying cancer-associated alterations using genomics. Here, we show that standard blood collection procedures rapidly change the transcriptional and posttranscriptional landscapes of hematopoietic cells, resulting in biased activation of specific biological pathways; up-regulation of pseudogenes, antisense RNAs, and unannotated coding isoforms; and RNA surveillance inhibition. Affected genes include common mutational targets and thousands of other genes participating in processes such as chromatin modification, RNA splicing, T-and B-cell activation, and NF-κB signaling. The majority of published leukemic transcriptomes exhibit signals of this incubation-induced dysregulation, explaining up to 40% of differences in gene expression and alternative splicing between leukemias and reference normal transcriptomes. The effects of sample processing are particularly evident in pan-cancer analyses. We provide biomarkers that detect prolonged incubation of individual samples and show that keeping blood on ice markedly reduces changes to the transcriptome. In addition to highlighting the potentially confounding effects of technical artifacts in cancer genomics data, our study emphasizes the need to survey the diversity of normal as well as neoplastic cells when characterizing tumors. leukemia | RNA splicing | nonsense-mediated decay | batch effect

Coordinated mis-splicing of TMEM14C and ABCB7 causes ring sideroblast formation in SF3B1-mutant myelodysplastic syndrome

F3B1 splicing factor mutations are near-universally found in myelodysplastic syndromes (MDS) with ring sideroblasts, a clonal hematopoietic disorder characterized by abnormal erythroid cells with iron-loaded mitochondria. Despite this remarkably strong genotype-to-phenotype correlation, the mechanism by which mutant SF3B1 dysregulates iron metabolism to cause ring sideroblasts (RS) remains unclear due to an absence of physiological models of RS formation. Here, we report an induced pluripotent stem cell (iPSC) model of SF3B1-mutant MDS that for the first time recapitulates robust RS formation during in vitro erythroid differentiation. Mutant SF3B1 induces mis-splicing of ~100 genes throughout erythroid differentiation, including proposed RS driver genes TMEM14C, PPOX, and ABCB7. All three mis-splicing events reduce protein expression, notably occurring via 5' UTR alteration and reduced translation efficiency for TMEM14C. Functional rescue of TMEM14C and ABCB7, but not the non-rate-limiting enzyme PPOX, markedly decreased RS, and their combined rescue nearly abolished RS formation. Our study demonstrates that coordinated mis-splicing of mitochondrial transporters TMEM14C and ABCB7 by mutant SF3B1 sequesters iron in mitochondria, causing ring sideroblast formation

SRSF2 Mutations Contribute to Myelodysplasia by Mutant-Specific Effects on Exon Recognition.

Mutations affecting spliceosomal proteins are the most common mutations in patients with myelodysplastic syndromes (MDS), but their role in MDS pathogenesis has not been delineated. Here we report that mutations affecting the splicing factor SRSF2 directly impair hematopoietic differentiation in vivo, which is not due to SRSF2 loss of function. By contrast, SRSF2 mutations alter SRSF2's normal sequence-specific RNA binding activity, thereby altering the recognition of specific exonic splicing enhancer motifs to drive recurrent mis-splicing of key hematopoietic regulators. This includes SRSF2 mutation-dependent splicing of EZH2, which triggers nonsense-mediated decay, which, in turn, results in impaired hematopoietic differentiation. These data provide a mechanistic link between a mutant spliceosomal protein, alterations in the splicing of key regulators, and impaired hematopoiesis.E.K. is supported by the Worldwide Cancer Research Fund. A.R. was supported by the NIH/NHLBI (U01 HL099993), NIH/NIDDK (K08 DK082783), the J.P. McCarthy Foundation, and the Storb Foundation. S.H. and O.A.-W. are supported by grants from the Edward P. Evans Foundation. S.H. was supported by Yale Comprehensive Cancer Center institutional funds. R.K.B. was supported by the Hartwell Innovation Fund, Damon Runyon Cancer Research Foundation (DFS 04-12), Ellison Medical Foundation (AG-NS-1030-13), NIH/NIDDK (R56 DK103854), NIH/NCI recruitment support (P30 CA015704), and Fred Hutchinson Cancer Research Center institutional funds. J.O.I. was supported by an NIH/NCI training grant (T32 CA009657) and NIH/NIDDK pilot study (P30 DK056465). C.L. is supported by a career development award grant from the Leukemia and Lymphoma Society and an ATIP-Avenir grant from the French government. O.A.-W. is supported by an NIH K08 clinical investigator award (1K08CA160647-01), a Department of Defense Postdoctoral Fellow Award in Bone Marrow Failure Research (W81XWH-12-1-0041), the Josie Robertson Investigator Program, and a Damon Runyon Clinical Investigator Award with support from the Evans Foundation. F.H.-T.A. acknowledges support from the NCCR RNA and Disease funded by the Swiss National Science Foundation and the SNF Sinergia CRSII3_127454. Y.L. and Y.M. were supported by NIH/NIGMS grant R01 GM102869 and Senior Research Fellowship Grant 101908/Z/13/Z (to Y.M.) from the Wellcome Trust. J.D. acknowledges assistance from Dr. Nezih Cereb, HistoGenetics (Ossining, NY).This is the author accepted manuscript. The final version is available from Elsevier via http://dx.doi.org/10.1016/j.ccell.2015.04.00