Skip to main content

Thank you for visiting You are using a browser version with limited support for CSS. To obtain the best experience, we recommend you use a more up to date browser (or turn off compatibility mode in Internet Explorer). In the meantime, to ensure continued support, we are displaying the site without styles and JavaScript.

Mechanisms of alternative splicing regulation: insights from molecular and genomics approaches

This article has been updated

Key Points

  • Alternative splicing is a crucial mechanism for gene regulation and for generating proteomic diversity, which allows individual genes to generate multiple mature mRNA isoforms that can be translated into functionally different proteins. Alternative splicing can be regulated at different stages of spliceosome assembly and by different mechanisms.

  • Splice site recognition of alternative exons is frequently regulated by cooperative interactions between factors such as SR (Ser–Arg) proteins and heterogeneous nuclear ribonucleoprotein particles (hnRNPs), which have lower affinities and sequence specificities. Splice site selection is also influenced by the secondary structure of mRNAs.

  • Two models have been proposed to explain the role of RNA polymerase II in alternative splicing regulation: a recruitment model and a kinetic model, and the two models are not mutually exclusive.

  • Alternative splicing, including tissue-specific alternative splicing, is an extremely common regulatory mechanism. However, the number of known sequence-specific alternative splicing factors (<50) is much smaller than that of sequence-specific transcription factors (2,500). Although more specific splicing factors will undoubtedly be discovered, this disparity suggests important differences in the pathways regulating transcription and splicing.

  • Core spliceosomal proteins can also regulate tissue-specific alternative splicing. This may reflect differential sensitivity of alternative exons to these factors and/or differential accumulation of the factors in different tissues.

  • Post-translational modifications of splicing factors enable cells to switch between alternative splicing isoforms rapidly after environmental stimuli. Phosphorylation can change the intracellular localization of splicing factor, protein–protein and protein–RNA interactions and even intrinsic splicing factor activity.


Alternative splicing of mRNA precursors provides an important means of genetic control and is a crucial step in the expression of most genes. Alternative splicing markedly affects human development, and its misregulation underlies many human diseases. Although the mechanisms of alternative splicing have been studied extensively, until the past few years we had not begun to realize fully the diversity and complexity of alternative splicing regulation by an intricate protein–RNA network. Great progress has been made by studying individual transcripts and through genome-wide approaches, which together provide a better picture of the mechanistic regulation of alternative pre-mRNA splicing.

Access options

Rent or Buy article

Get time limited or full article access on ReadCube.


All prices are NET prices.

Figure 1: Mechanisms of alternative splicing by splice site selection.
Figure 2: Phosphorylation switches the general splicing repressor SRp38 into a sequence-specific activator.
Figure 3: Mechanisms of alternative splicing regulation at the transition from exon definition to intron definition.

Change history

  • 12 October 2009

    In the version of this article initially published online the binding sequences for hnRNP G and hnRNP Q in Table 1 were incorrect. This error has been corrected for the print, HTML and PDF versions of the article.


  1. 1

    Black, D. L. Mechanisms of alternative pre-messenger RNA splicing. Annu. Rev. Biochem. 72, 291–336 (2003).

    CAS  PubMed  Google Scholar 

  2. 2

    Wahl, M. C., Will, C. L. & Luhrmann, R. The spliceosome: design principles of a dynamic RNP machine. Cell 136, 701–718 (2009).

    CAS  PubMed  Google Scholar 

  3. 3

    Berglund, J. A., Chua, K., Abovich, N., Reed, R. & Rosbash, M. The splicing factor BBP interacts specifically with the pre-mRNA branchpoint sequence UACUAAC. Cell 89, 781–787 (1997).

    CAS  Google Scholar 

  4. 4

    Zamore, P. D. & Green, M. R. Identification, purification, and biochemical characterization of U2 small nuclear ribonucleoprotein auxiliary factor. Proc. Natl Acad. Sci. USA 86, 9243–9247 (1989).

    CAS  Google Scholar 

  5. 5

    Nelson, K. K. & Green, M. R. Mammalian U2 snRNP has a sequence-specific RNA-binding activity. Genes Dev. 3, 1562–1571 (1989).

    CAS  Google Scholar 

  6. 6

    Graveley, B. R. Sorting out the complexity of SR protein functions. RNA 6, 1197–1211 (2000).

    CAS  PubMed  PubMed Central  Google Scholar 

  7. 7

    Tacke, R. & Manley, J. L. Determinants of SR protein specificity. Curr. Opin. Cell Biol. 11, 358–362 (1999).

    CAS  Google Scholar 

  8. 8

    Long, J. C. & Caceres, J. F. The SR protein family of splicing factors: master regulators of gene expression. Biochem. J. 417, 15–27 (2009).

    CAS  Google Scholar 

  9. 9

    Smith, C. W. & Valcarcel, J. Alternative pre-mRNA splicing: the logic of combinatorial control. Trends Biochem. Sci. 25, 381–388 (2000).

    CAS  Google Scholar 

  10. 10

    Dreyfuss, G., Kim, V. N. & Kataoka, N. Messenger-RNA-binding proteins and the messages they carry. Nature Rev. Mol. Cell Biol. 3, 195–205 (2002).

    CAS  Google Scholar 

  11. 11

    Ule, J. et al. An RNA map predicting Nova-dependent splicing regulation. Nature 444, 580–586 (2006).

    CAS  PubMed  Google Scholar 

  12. 12

    Hui, J. et al. Intronic CA-repeat and CA-rich elements: a new class of regulators of mammalian alternative splicing. EMBO J. 24, 1988–1998 (2005).

    CAS  PubMed  PubMed Central  Google Scholar 

  13. 13

    Yeo, G. W. et al. An RNA code for the FOX2 splicing regulator revealed by mapping RNA-protein interactions in stem cells. Nature Struct. Mol. Biol. 16, 130–137 (2009).

    CAS  Google Scholar 

  14. 14

    Mauger, D. M., Lin, C. & Garcia-Blanco, M. A. hnRNP H and hnRNP F complex with Fox2 to silence fibroblast growth factor receptor 2 exon IIIc. Mol. Cell. Biol. 28, 5403–5419 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  15. 15

    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).

    CAS  Google Scholar 

  16. 16

    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). This study shows that PTB inhibits SRC exon N1 inclusion by preventing the transition from an exon-definition to an intron-definition complex and analyses the protein composition of different complexes.

    CAS  Google Scholar 

  17. 17

    Lallena, M. J., Chalmers, K. J., Llamazares, S., Lamond, A. I. & Valcarcel, J. Splicing regulation at the second catalytic step by Sex-lethal involves 3′ splice site recognition by SPF45. Cell 109, 285–296 (2002).

    CAS  Google Scholar 

  18. 18

    Batsche, E., Yaniv, M. & Muchardt, C. The human SWI/SNF subunit Brm is a regulator of alternative splicing. Nature Struct. Mol. Biol. 13, 22–29 (2006). This study shows that BRM promotes the inclusion of variable exons of CD44 pre-mRNA by stalling RNAP II at the variable exon-containing region of the CD44 gene. It also shows that BRM interacts with splicing factor SAM68.

    CAS  Google Scholar 

  19. 19

    de la Mata, M. & Kornblihtt, A. R. RNA polymerase II C-terminal domain mediates regulation of alternative splicing by SRp20. Nature Struct. Mol. Biol. 13, 973–980 (2006).

    CAS  Google Scholar 

  20. 20

    Sims, R. J. 3rd et al. Recognition of trimethylated histone H3 lysine 4 facilitates the recruitment of transcription postinitiation factors and pre-mRNA splicing. Mol. Cell 28, 665–676 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  21. 21

    Lin, S., Coutinho-Mansfield, G., Wang, D., Pandit, S. & Fu, X. D. The splicing factor SC35 has an active role in transcriptional elongation. Nature Struct. Mol. Biol. 15, 819–826 (2008).

    CAS  Google Scholar 

  22. 22

    Moldon, A. et al. Promoter-driven splicing regulation in fission yeast. Nature 455, 997–1000 (2008).

    CAS  Google Scholar 

  23. 23

    Graveley, B. R. Alternative splicing: increasing diversity in the proteomic world. Trends Genet. 17, 100–107 (2001).

    CAS  Google Scholar 

  24. 24

    Blencowe, B. J. & Graveley, B. R. (eds) Alternative Splicing in the Postgenomic Era. (Springer, the Netherlands, 2007).

    Google Scholar 

  25. 25

    Grabowski, P. J. & Black, D. L. Alternative RNA splicing in the nervous system. Prog. Neurobiol. 65, 289–308 (2001).

    CAS  Google Scholar 

  26. 26

    Park, J. W., Parisky, K., Celotto, A. M., Reenan, R. A. & Graveley, B. R. Identification of alternative splicing regulators by RNA interference in Drosophila. Proc. Natl Acad. Sci. USA 101, 15974–15979 (2004).

    CAS  Google Scholar 

  27. 27

    Zhang, Z. et al. SMN deficiency causes tissue-specific perturbations in the repertoire of snRNAs and widespread defects in splicing. Cell 133, 585–600 (2008). This paper shows that SMN deficiency regulates snRNP levels in a tissue-specific manner, and this was reflected in altered alternative splicing patterns in different mouse tissues.

    CAS  PubMed  PubMed Central  Google Scholar 

  28. 28

    Blencowe, B. J. Alternative splicing: new insights from global analyses. Cell 126, 37–47 (2006).

    CAS  Google Scholar 

  29. 29

    Wang, E. T. et al. Alternative isoform regulation in human tissue transcriptomes. Nature 456, 470–476 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  30. 30

    Pan, Q., Shai, O., Lee, L. J., Frey, B. J. & Blencowe, B. J. Deep surveying of alternative splicing complexity in the human transcriptome by high-throughput sequencing. Nature Genet. 40, 1413–1415 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  31. 31

    Castle, J. C. et al. Expression of 24,426 human alternative splicing events and predicted cis regulation in 48 tissues and cell lines. Nature Genet. 40, 1416–1425 (2008). A transcriptome study that analyses alternative splicing events from 48 tissues and identifies tissue-specific regulatory motifs and cognate binding proteins.

    CAS  PubMed  Google Scholar 

  32. 32

    Sultan, M. et al. A global view of gene activity and alternative splicing by deep sequencing of the human transcriptome. Science 321, 956–960 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  33. 33

    Sterner, D. A., Carlo, T. & Berget, S. M. Architectural limits on split genes. Proc. Natl Acad. Sci. USA 93, 15081–15085 (1996).

    CAS  Google Scholar 

  34. 34

    Berget, S. M. Exon recognition in vertebrate splicing. J. Biol. Chem. 270, 2411–2414 (1995).

    CAS  Google Scholar 

  35. 35

    Lim, S. R. & Hertel, K. J. Commitment to splice site pairing coincides with A complex formation. Mol. Cell 15, 477–483 (2004).

    CAS  Google Scholar 

  36. 36

    Kotlajich, M. V., Crabb, T. L. & Hertel, K. J. Spliceosome assembly pathways for different types of alternative splicing converge during commitment to splice site pairing in the A complex. Mol. Cell. Biol. 29, 1072–1082 (2009). This study shows that the commitment to splicing of some alternative exons occurs during splice site pairing in the A complex and that ATP hydrolysis is required for splice site paring, thereby locking splice sites into a splicing pattern after U2 snRNP binding to the branch site.

    CAS  Google Scholar 

  37. 37

    Bourgeois, C. F., Popielarz, M., Hildwein, G. & Stevenin, J. Identification of a bidirectional splicing enhancer: differential involvement of SR proteins in 5′ or 3′ splice site activation. Mol. Cell. Biol. 19, 7347–7356 (1999).

    CAS  PubMed  PubMed Central  Google Scholar 

  38. 38

    Zuo, P. & Maniatis, T. The splicing factor U2AF35 mediates critical protein–protein interactions in constitutive and enhancer-dependent splicing. Genes Dev. 10, 1356–1368 (1996).

    CAS  Google Scholar 

  39. 39

    Feng, Y., Chen, M. & Manley, J. L. Phosphorylation switches the general splicing repressor SRp38 to a sequence-specific activator. Nature Struct. Mol. Biol. 15, 1040–1048 (2008). This paper shows that phosphorylation switches SRp38 from a general repressor to a sequence-specific activator that functions by recruiting and stabilizing U1 and U2 snRNP at splice sites.

    CAS  Google Scholar 

  40. 40

    Graveley, B. R., Hertel, K. J. & Maniatis, T. The role of U2AF35 and U2AF65 in enhancer-dependent splicing. RNA 7, 806–818 (2001).

    CAS  PubMed  PubMed Central  Google Scholar 

  41. 41

    Kohtz, J. D. et al. Protein–protein interactions and 5′-splice-site recognition in mammalian mRNA precursors. Nature 368, 119–124 (1994).

    CAS  Google Scholar 

  42. 42

    Wu, J. Y. & Maniatis, T. Specific interactions between proteins implicated in splice site selection and regulated alternative splicing. Cell 75, 1061–1070 (1993).

    CAS  Google Scholar 

  43. 43

    Xiao, S. H. & Manley, J. L. Phosphorylation of the ASF/SF2 RS domain affects both protein-protein and protein-RNA interactions and is necessary for splicing. Genes Dev. 11, 334–344 (1997).

    CAS  Google Scholar 

  44. 44

    Pacheco, T. R., Coelho, M. B., Desterro, J. M., Mollet, I. & Carmo-Fonseca, M. In vivo requirement of the small subunit of U2AF for recognition of a weak 3′ splice site. Mol. Cell. Biol. 26, 8183–90 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  45. 45

    Longman, D. et al. Multiple interactions between SRm160 and SR family proteins in enhancer-dependent splicing and development of C. elegans. Curr. Biol. 11, 1923–1933 (2001).

    CAS  Google Scholar 

  46. 46

    Blencowe, B. J. et al. The SRm160/300 splicing coactivator subunits. RNA 6, 111–120 (2000).

    CAS  PubMed  PubMed Central  Google Scholar 

  47. 47

    Tacke, R. & Manley, J. L. Functions of SR and Tra2 proteins in pre-mRNA splicing regulation. Proc. Soc. Exp. Biol. Med. 220, 59–63 (1999).

    CAS  Google Scholar 

  48. 48

    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).

    CAS  Google Scholar 

  49. 49

    Forch, P., Puig, O., Martinez, C., Seraphin, B. & Valcarcel, J. The splicing regulator TIA-1 interacts with U1-C to promote U1 snRNP recruitment to 5′ splice sites. EMBO J. 21, 6882–6892 (2002).

    PubMed  PubMed Central  Google Scholar 

  50. 50

    Tisserant, A. & Konig, H. Signal-regulated pre-mRNA occupancy by the general splicing factor U2AF. PLoS ONE 3, e1418 (2008).

    PubMed  PubMed Central  Google Scholar 

  51. 51

    Yang, L., Embree, L. J., Tsai, S. & Hickstein, D. D. Oncoprotein TLS interacts with serine-arginine proteins involved in RNA splicing. J. Biol. Chem. 273, 27761–27764 (1998).

    CAS  Google Scholar 

  52. 52

    Komatsu, M., Kominami, E., Arahata, K. & Tsukahara, T. Cloning and characterization of two neural-salient serine/arginine-rich (NSSR) proteins involved in the regulation of alternative splicing in neurones. Genes Cells 4, 593–606 (1999).

    CAS  Google Scholar 

  53. 53

    Cowper, A. E., Caceres, J. F., Mayeda, A. & Screaton, G. R. Serine-arginine (SR) protein-like factors that antagonize authentic SR proteins and regulate alternative splicing. J. Biol. Chem. 276, 48908–48914 (2001).

    CAS  Google Scholar 

  54. 54

    Shin, C. & Manley, J. L. The SR protein SRp38 represses splicing in M phase cells. Cell 111, 407–417 (2002).

    CAS  Google Scholar 

  55. 55

    Shin, C., Feng, Y. & Manley, J. L. Dephosphorylated SRp38 acts as a splicing repressor in response to heat shock. Nature 427, 553–558 (2004).

    CAS  Google Scholar 

  56. 56

    Krainer, A. R., Conway, G. C. & Kozak, D. Purification and characterization of pre-mRNA splicing factor SF2 from HeLa cells. Genes Dev. 4, 1158–1171 (1990).

    CAS  Google Scholar 

  57. 57

    Singh, R., Valcarcel, J. & Green, M. R. Distinct binding specificities and functions of higher eukaryotic polypyrimidine tract-binding proteins. Science 268, 1173–1176 (1995).

    CAS  Google Scholar 

  58. 58

    Spellman, R. & Smith, C. W. Novel modes of splicing repression by PTB. Trends Biochem. Sci. 31, 73–76 (2006).

    CAS  PubMed  Google Scholar 

  59. 59

    Sauliere, J., Sureau, A., Expert-Bezancon, A. & Marie, J. The polypyrimidine tract binding protein (PTB) represses splicing of exon 6B from the β-tropomyosin pre-mRNA by directly interfering with the binding of the U2AF65 subunit. Mol. Cell. Biol. 26, 8755–8769 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  60. 60

    Tange, T. O., Damgaard, C. K., Guth, S., Valcarcel, J. & Kjems, J. The hnRNP A1 protein regulates HIV-1 tat splicing via a novel intron silencer element. EMBO J. 20, 5748–5758 (2001).

    CAS  PubMed  PubMed Central  Google Scholar 

  61. 61

    Zhou, H. L. & Lou, H. Repression of prespliceosome complex formation at two distinct steps by Fox-1/Fox-2 proteins. Mol. Cell. Biol. 28, 5507–5516 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  62. 62

    Zhu, H., Hinman, M. N., Hasman, R. A., Mehta, P. & Lou, H. Regulation of neuron-specific alternative splicing of neurofibromatosis type 1 pre-mRNA. Mol. Cell. Biol. 28, 1240–1251 (2008).

    CAS  Google Scholar 

  63. 63

    Kashima, T. & Manley, J. L. A negative element in SMN2 exon 7 inhibits splicing in spinal muscular atrophy. Nature Genet. 34, 460–463 (2003).

    CAS  PubMed  Google Scholar 

  64. 64

    Martins de Araujo, M., Bonnal, S., Hastings, M. L., Krainer, A. R. & Valcarcel, J. Differential 3′ splice site recognition of SMN1 and SMN2 transcripts by U2AF and U2 snRNP. RNA 15, 515–523 (2009).

    PubMed  PubMed Central  Google Scholar 

  65. 65

    Sharma, S., Falick, A. M. & Black, D. L. Polypyrimidine tract binding protein blocks the 5′ splice site-dependent assembly of U2AF and the prespliceosomal E complex. Mol. Cell 19, 485–496 (2005).

    CAS  PubMed  PubMed Central  Google Scholar 

  66. 66

    Damgaard, C. K., Tange, T. O. & Kjems, J. hnRNP A1 controls HIV-1 mRNA splicing through cooperative binding to intron and exon splicing silencers in the context of a conserved secondary structure. RNA 8, 1401–1415 (2002).

    CAS  PubMed  PubMed Central  Google Scholar 

  67. 67

    Nasim, F. U., Hutchison, S., Cordeau, M. & Chabot, B. High-affinity hnRNP A1 binding sites and duplex-forming inverted repeats have similar effects on 5′ splice site selection in support of a common looping out and repression mechanism. RNA 8, 1078–1089 (2002).

    CAS  PubMed  PubMed Central  Google Scholar 

  68. 68

    Hutchison, S., LeBel, C., Blanchette, M. & Chabot, B. Distinct sets of adjacent heterogeneous nuclear ribonucleoprotein (hnRNP) A1/A2 binding sites control 5′ splice site selection in the hnRNP A1 mRNA precursor. J. Biol. Chem. 277, 29745–29752 (2002).

    CAS  Google Scholar 

  69. 69

    Kashima, T., Rao, N., David, C. J. & Manley, J. L. hnRNP A1 functions with specificity in repression of SMN2 exon 7 splicing. Hum. Mol. Genet. 16, 3149–3159 (2007).

    CAS  Google Scholar 

  70. 70

    Kashima, T., Rao, N. & Manley, J. L. An intronic element contributes to splicing repression in spinal muscular atrophy. Proc. Natl Acad. Sci. USA 104, 3426–3431 (2007).

    CAS  Google Scholar 

  71. 71

    Chou, M. Y., Underwood, J. G., Nikolic, J., Luu, M. H. & Black, D. L. Multisite RNA binding and release of polypyrimidine tract binding protein during the regulation of c-src neural-specific splicing. Mol. Cell 5, 949–957 (2000).

    CAS  Google Scholar 

  72. 72

    Mayeda, A., Helfman, D. M. & Krainer, A. R. Modulation of exon skipping and inclusion by heterogeneous nuclear ribonucleoprotein A1 and pre-mRNA splicing factor SF2/ASF. Mol. Cell. Biol. 13, 2993–3001 (1993).

    CAS  PubMed  PubMed Central  Google Scholar 

  73. 73

    Zahler, A. M., Damgaard, C. K., Kjems, J. & Caputi, M. SC35 and heterogeneous nuclear ribonucleoprotein A/B proteins bind to a juxtaposed exonic splicing enhancer/exonic splicing silencer element to regulate HIV-1 tat exon 2 splicing. J. Biol. Chem. 279, 10077–10084 (2004).

    CAS  Google Scholar 

  74. 74

    Zhu, J. & Krainer, A. R. Pre-mRNA splicing in the absence of an SR protein RS domain. Genes Dev. 14, 3166–3178 (2000).

    CAS  PubMed  PubMed Central  Google Scholar 

  75. 75

    Crawford, J. B. & Patton, J. G. Activation of α-tropomyosin exon 2 is regulated by the SR protein 9G8 and heterogeneous nuclear ribonucleoproteins H and F. Mol. Cell. Biol. 26, 8791–802 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  76. 76

    Expert-Bezancon, A. et al. hnRNP A1 and the SR proteins ASF/SF2 and SC35 have antagonistic functions in splicing of β-tropomyosin exon 6B. J. Biol. Chem. 279, 38249–59 (2004).

    CAS  Google Scholar 

  77. 77

    Charlet, B. N., Logan, P., Singh, G. & Cooper, T. A. Dynamic antagonism between ETR-3 and PTB regulates cell type-specific alternative splicing. Mol. Cell 9, 649–658 (2002).

    Google Scholar 

  78. 78

    Blanchette, M. et al. Genome-wide analysis of alternative pre-mRNA splicing and RNA-binding specificities of the Drosophila hnRNP A/B family members. Mol. Cell 33, 438–449 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  79. 79

    Hung, L. H. et al. Diverse roles of hnRNP L in mammalian mRNA processing: a combined microarray and RNAi analysis. RNA 14, 284–296 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  80. 80

    Licatalosi, D. D. et al. HITS-CLIP yields genome-wide insights into brain alternative RNA processing. Nature 456, 464–469 (2008). This paper introduces a method that combines CLIP and high-throughput sequencing to identify targets of NOVA proteins.

    CAS  PubMed  PubMed Central  Google Scholar 

  81. 81

    Dredge, B. K., Stefani, G., Engelhard, C. C. & Darnell, R. B. Nova autoregulation reveals dual functions in neuronal splicing. EMBO J. 24, 1608–1620 (2005).

    CAS  PubMed  PubMed Central  Google Scholar 

  82. 82

    Martinez-Contreras, R. et al. Intronic binding sites for hnRNP A/B and hnRNP F/H proteins stimulate pre-mRNA splicing. PLoS Biol. 4, e21 (2006).

    PubMed  PubMed Central  Google Scholar 

  83. 83

    Wang, Z. & Burge, C. B. Splicing regulation: from a parts list of regulatory elements to an integrated splicing code. RNA 14, 802–813 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  84. 84

    Dredge, B. K. & Darnell, R. B. Nova regulates GABAA receptor γ2 alternative splicing via a distal downstream UCAU-rich intronic splicing enhancer. Mol. Cell. Biol. 23, 4687–4700 (2003).

    CAS  PubMed  PubMed Central  Google Scholar 

  85. 85

    Schaub, M. C., Lopez, S. R. & Caputi, M. Members of the heterogeneous nuclear ribonucleoprotein H family activate splicing of an HIV-1 splicing substrate by promoting formation of ATP-dependent spliceosomal complexes. J. Biol. Chem. 282, 13617–13626 (2007).

    CAS  Google Scholar 

  86. 86

    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).

    CAS  Google Scholar 

  87. 87

    Sanford, J. R. et al. Identification of nuclear and cytoplasmic mRNA targets for the shuttling protein SF2/ASF. PLoS ONE 3, e3369 (2008).

    PubMed  PubMed Central  Google Scholar 

  88. 88

    Sanford, J. R. et al. Splicing factor SFRS1 recognizes a functionally diverse landscape of RNA transcripts. Genome Res. 19, 381–394 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  89. 89

    Graveley, B. R. Mutually exclusive splicing of the insect Dscam pre-mRNA directed by competing intronic RNA secondary structures. Cell 123, 65–73 (2005).

    CAS  PubMed  PubMed Central  Google Scholar 

  90. 90

    Olson, S. et al. A regulator of Dscam mutually exclusive splicing fidelity. Nature Struct. Mol. Biol. 14, 1134–1140 (2007).

    CAS  Google Scholar 

  91. 91

    Grover, A. et al. 5′ splice site mutations in tau associated with the inherited dementia FTDP-17 affect a stem-loop structure that regulates alternative splicing of exon 10. J. Biol. Chem. 274, 15134–15143 (1999).

    CAS  Google Scholar 

  92. 92

    Hiller, M., Zhang, Z., Backofen, R. & Stamm, S. Pre-mRNA secondary structures influence exon recognition. PLoS Genet. 3, e204 (2007).

    PubMed  PubMed Central  Google Scholar 

  93. 93

    Camats, M., Guil, S., Kokolo, M. & Bach-Elias, M. P68 RNA helicase (DDX5) alters activity of cis- and trans-acting factors of the alternative splicing of H-Ras. PLoS ONE 3, e2926 (2008).

    PubMed  PubMed Central  Google Scholar 

  94. 94

    Libri, D., Balvay, L. & Fiszman, M. Y. In vivo splicing of the beta tropomyosin pre-mRNA: a role for branch point and donor site competition. Mol. Cell. Biol. 12, 3204–3215 (1992).

    CAS  PubMed  PubMed Central  Google Scholar 

  95. 95

    Henkin, T. M. Riboswitch RNAs: using RNA to sense cellular metabolism. Genes Dev. 22, 3383–3390 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  96. 96

    Cheah, M. T., Wachter, A., Sudarsan, N. & Breaker, R. R. Control of alternative RNA splicing and gene expression by eukaryotic riboswitches. Nature 447, 497–500 (2007).

    CAS  PubMed  Google Scholar 

  97. 97

    Kishore, S. & Stamm, S. Regulation of alternative splicing by snoRNAs. Cold Spring Harb. Symp. Quant. Biol. 71, 329–334 (2006).

    CAS  Google Scholar 

  98. 98

    Kishore, S. & Stamm, S. The snoRNA HBII-52 regulates alternative splicing of the serotonin receptor 2C. Science 311, 230–232 (2006).

    CAS  PubMed  Google Scholar 

  99. 99

    Yu, Y. et al. Dynamic regulation of alternative splicing by silencers that modulate 5′ splice site competition. Cell 135, 1224–1236 (2008). The authors screen for splicing silencers that favour the inclusion of a distal 5′ splice site and provide evidence that the silencers work by changing the conformation of the pre-mRNA–U1 snRNP complex and promoting pairing of the distal U1 snRNP with U2 snRNP.

    CAS  PubMed  PubMed Central  Google Scholar 

  100. 100

    Makeyev, E. V., Zhang, J., Carrasco, M. A. & Maniatis, T. The microRNA miR-124 promotes neuronal differentiation by triggering brain-specific alternative pre-mRNA splicing. Mol. Cell 27, 435–448 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  101. 101

    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). Using siRNA and microarray analysis, the authors show that the PTB-to-nPTB switch provides a post-transcriptional mechanism that is important for programming neuronal differentiation.

    CAS  PubMed  PubMed Central  Google Scholar 

  102. 102

    Coutinho-Mansfield, G. C., Xue, Y., Zhang, Y. & Fu, X. D. PTB/nPTB switch: a post-transcriptional mechanism for programming neuronal differentiation. Genes Dev. 21, 1573–1577 (2007).

    CAS  Google Scholar 

  103. 103

    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).

    CAS  PubMed  PubMed Central  Google Scholar 

  104. 104

    Ohi, M. D. et al. Structural and functional analysis of essential pre-mRNA splicing factor Prp19p. Mol. Cell. Biol. 25, 451–460 (2005).

    CAS  PubMed  PubMed Central  Google Scholar 

  105. 105

    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).

    CAS  Google Scholar 

  106. 106

    Eldridge, A. G., Li, Y., Sharp, P. A. & Blencowe, B. J. The SRm160/300 splicing coactivator is required for exon-enhancer function. Proc. Natl Acad. Sci. USA 96, 6125–6130 (1999).

    CAS  Google Scholar 

  107. 107

    Edamatsu, H., Kaziro, Y. & Itoh, H. LUCA15, a putative tumour suppressor gene encoding an RNA-binding nuclear protein, is down-regulated in ras-transformed Rat-1 cells. Genes Cells 5, 849–858 (2000).

    CAS  Google Scholar 

  108. 108

    Mourtada-Maarabouni, M., Sutherland, L. C. & Williams, G. T. Candidate tumour suppressor LUCA-15 can regulate multiple apoptotic pathways. Apoptosis 7, 421–432 (2002).

    CAS  Google Scholar 

  109. 109

    Bonnal, S. et al. RBM5/Luca-15/H37 regulates Fas alternative splice site pairing after exon definition. Mol. Cell 32, 81–95 (2008).

    CAS  Google Scholar 

  110. 110

    Kornblihtt, A. R. Chromatin, transcript elongation and alternative splicing. Nature Struct. Mol. Biol. 13, 5–7 (2006).

    CAS  Google Scholar 

  111. 111

    Das, R. et al. SR proteins function in coupling RNAP II transcription to pre-mRNA splicing. Mol. Cell 26, 867–881 (2007).

    CAS  Google Scholar 

  112. 112

    Auboeuf, D. et al. Differential recruitment of nuclear receptor coactivators may determine alternative RNA splice site choice in target genes. Proc. Natl Acad. Sci. USA 101, 2270–2274 (2004).

    CAS  Google Scholar 

  113. 113

    Cramer, P. et al. Coupling of transcription with alternative splicing: RNA pol II promoters modulate SF2/ASF and 9G8 effects on an exonic splicing enhancer. Mol. Cell 4, 251–258 (1999).

    CAS  Google Scholar 

  114. 114

    Monsalve, M. et al. Direct coupling of transcription and mRNA processing through the thermogenic coactivator PGC-1. Mol. Cell 6, 307–316 (2000).

    CAS  Google Scholar 

  115. 115

    de la Mata, M. et al. A slow RNA polymerase II affects alternative splicing in vivo. Mol. Cell 12, 525–532 (2003).

    CAS  Google Scholar 

  116. 116

    Munoz, M. J. et al. DNA damage regulates alternative splicing through inhibition of RNA polymerase II elongation. Cell 137, 708–720 (2009).

    CAS  Google Scholar 

  117. 117

    Phatnani, H. P. & Greenleaf, A. L. Phosphorylation and functions of the RNA polymerase II CTD. Genes Dev. 20, 2922–2936 (2006).

    CAS  PubMed  Google Scholar 

  118. 118

    Lin, P. S., Marshall, N. F. & Dahmus, M. E. CTD phosphatase: role in RNA polymerase II cycling and the regulation of transcript elongation. Prog. Nucleic Acid Res. Mol. Biol. 72, 333–365 (2002).

    CAS  Google Scholar 

  119. 119

    David, C. J. & Manley, J. L. The search for alternative splicing regulators: new approaches offer a path to a splicing code. Genes Dev. 22, 279–285 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  120. 120

    Ohno, G., Hagiwara, M. & Kuroyanagi, H. STAR family RNA-binding protein ASD-2 regulates developmental switching of mutually exclusive alternative splicing in vivo. Genes Dev. 22, 360–374 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  121. 121

    Li, Q., Lee, J. A. & Black, D. L. Neuronal regulation of alternative pre-mRNA splicing. Nature Rev. Neurosci. 8, 819–831 (2007).

    CAS  Google Scholar 

  122. 122

    Ule, J. et al. Nova regulates brain-specific splicing to shape the synapse. Nature Genet. 37, 844–852 (2005).

    CAS  Google Scholar 

  123. 123

    Perrone-Bizzozero, N. & Bolognani, F. Role of HuD and other RNA-binding proteins in neural development and plasticity. J. Neurosci. Res. 68, 121–126 (2002).

    CAS  Google Scholar 

  124. 124

    Zhu, H., Hasman, R. A., Barron, V. A., Luo, G. & Lou, H. A nuclear function of Hu proteins as neuron-specific alternative RNA processing regulators. Mol. Biol. Cell 17, 5105–5114 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  125. 125

    Soller, M., Li, M. & Haussmann, I. U. Regulation of the ELAV target ewg: insights from an evolutionary perspective. Biochem. Soc. Trans. 36, 502–504 (2008).

    CAS  Google Scholar 

  126. 126

    McKee, A. E. et al. A genome-wide in situ hybridization map of RNA-binding proteins reveals anatomically restricted expression in the developing mouse brain. BMC Dev. Biol. 5, 14 (2005).

    PubMed  PubMed Central  Google Scholar 

  127. 127

    Yang, Y. Y., Yin, G. L. & Darnell, R. B. The neuronal RNA-binding protein Nova-2 is implicated as the autoantigen targeted in POMA patients with dementia. Proc. Natl Acad. Sci. USA 95, 13254–13259 (1998).

    CAS  Google Scholar 

  128. 128

    Warzecha, C. C., Sato, T. K., Nabet, B., Hogenesch, J. B. & Carstens, R. P. ESRP1 and ESRP2 are epithelial cell-type-specific regulators of FGFR2 splicing. Mol. Cell 33, 591–601 (2009). The authors identify ESRP1 and ESRP2 as epithelial cell-specific alternative splicing factors and find that they regulate several epithelial cell-specific exons. They also show that ESRP1 and ESRP2 expression levels correlate with the splicing pattern change that is observed during the epithelial-to-mesenchymal cell transition.

    CAS  PubMed  PubMed Central  Google Scholar 

  129. 129

    Ding, J. H. et al. Dilated cardiomyopathy caused by tissue-specific ablation of SC35 in the heart. EMBO J. 23, 885–896 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

  130. 130

    Xu, X. et al. ASF/SF2-regulated CaMKIIδ alternative splicing temporally reprograms excitation-contraction coupling in cardiac muscle. Cell 120, 59–72 (2005).

    CAS  Google Scholar 

  131. 131

    Feng, Y. et al. SRp38 regulates alternative splicing and is required for Ca2+ handling in the embryonic heart. Dev. Cell 16, 528–538 (2009). This study shows that mice lacking SRp38 die perinatally and have multiple cardiac defects, and that the mRNA encoding cardiac triadin is a direct target of SRp38.

    CAS  PubMed  PubMed Central  Google Scholar 

  132. 132

    Grosso, A. R. et al. Tissue-specific splicing factor gene expression signatures. Nucleic Acids Res. 36, 4823–4832 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  133. 133

    Massiello, A., Roesser, J. R. & Chalfant, C. E. SAP155 binds to ceramide-responsive RNA cis-element 1 and regulates the alternative 5′ splice site selection of Bcl-x pre-mRNA. FASEB J. 20, 1680–1682 (2006).

    CAS  Google Scholar 

  134. 134

    Pacheco, T. R., Moita, L. F., Gomes, A. Q., Hacohen, N. & Carmo-Fonseca, M. RNA interference knockdown of hU2AF35 impairs cell cycle progression and modulates alternative splicing of Cdc25 transcripts. Mol. Biol. Cell 17, 4187–4199 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  135. 135

    Pleiss, J. A., Whitworth, G. B., Bergkessel, M. & Guthrie, C. Rapid, transcript-specific changes in splicing in response to environmental stress. Mol. Cell 27, 928–937 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  136. 136

    Monani, U. R. Spinal muscular atrophy: a deficiency in a ubiquitous protein; a motor neuron-specific disease. Neuron 48, 885–896 (2005).

    CAS  Google Scholar 

  137. 137

    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).

    PubMed  PubMed Central  Google Scholar 

  138. 138

    Shin, C. & Manley, J. L. Cell signalling and the control of pre-mRNA splicing. Nature Rev. Mol. Cell Biol. 5, 727–738 (2004).

    CAS  Google Scholar 

  139. 139

    Tarn, W. Y. Cellular signals modulate alternative splicing. J. Biomed. Sci. 14, 517–522 (2007).

    CAS  Google Scholar 

  140. 140

    Huang, Y., Yario, T. A. & Steitz, J. A. A molecular link between SR protein dephosphorylation and mRNA export. Proc. Natl Acad. Sci. USA 101, 9666–9670 (2004).

    CAS  Google Scholar 

  141. 141

    van der Houven van Oordt, W. et al. The MKK(3/6)-p38-signaling cascade alters the subcellular distribution of hnRNP A1 and modulates alternative splicing regulation. J. Cell Biol. 149, 307–316 (2000).

    CAS  PubMed  PubMed Central  Google Scholar 

  142. 142

    Habelhah, H. et al. ERK phosphorylation drives cytoplasmic accumulation of hnRNP-K and inhibition of mRNA translation. Nature Cell Biol. 3, 325–330 (2001).

    CAS  Google Scholar 

  143. 143

    Daoud, R. et al. Ischemia induces a translocation of the splicing factor tra2-beta 1 and changes alternative splicing patterns in the brain. J. Neurosci. 22, 5889–5899 (2002).

    CAS  Google Scholar 

  144. 144

    Guil, S., Long, J. C. & Caceres, J. F. hnRNP A1 relocalization to the stress granules reflects a role in the stress response. Mol. Cell. Biol. 26, 5744–5758 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  145. 145

    Huang, C. J., Tang, Z., Lin, R. J. & Tucker, P. W. Phosphorylation by SR kinases regulates the binding of PTB-associated splicing factor (PSF) to the pre-mRNA polypyrimidine tract. FEBS Lett. 581, 223–232 (2007).

    CAS  Google Scholar 

  146. 146

    Tacke, R., Chen, Y. & Manley, J. L. Sequence-specific RNA binding by an SR protein requires RS domain phosphorylation: creation of an SRp40-specific splicing enhancer. Proc. Natl Acad. Sci. USA 94, 1148–1153 (1997).

    CAS  Google Scholar 

  147. 147

    Izquierdo, J. M. & Valcarcel, J. Fas-activated serine/threonine kinase (FAST K) synergizes with TIA-1/TIAR proteins to regulate Fas alternative splicing. J. Biol. Chem. 282, 1539–1543 (2007).

    CAS  Google Scholar 

  148. 148

    Paronetto, M. P., Achsel, T., Massiello, A., Chalfant, C. E. & Sette, C. The RNA-binding protein Sam68 modulates the alternative splicing of Bcl-x. J. Cell Biol. 176, 929–939 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  149. 149

    Ma, S., Liu, G., Sun, Y. & Xie, J. Relocalization of the polypyrimidine tract-binding protein during PKA-induced neurite growth. Biochim. Biophys. Acta 1773, 912–923 (2007).

    CAS  Google Scholar 

  150. 150

    Xie, J., Lee, J. A., Kress, T. L., Mowry, K. L. & Black, D. L. Protein kinase A phosphorylation modulates transport of the polypyrimidine tract-binding protein. Proc. Natl Acad. Sci. USA 100, 8776–8781 (2003).

    CAS  Google Scholar 

  151. 151

    Shi, Y. & Manley, J. L. A complex signaling pathway regulates SRp38 phosphorylation and pre-mRNA splicing in response to heat shock. Mol. Cell 28, 79–90 (2007).

    CAS  Google Scholar 

  152. 152

    Van Eynde, A. et al. Molecular cloning of NIPP-1, a nuclear inhibitor of protein phosphatase-1, reveals homology with polypeptides involved in RNA processing. J. Biol. Chem. 270, 28068–28074 (1995).

    CAS  Google Scholar 

  153. 153

    Cohen, P. T. Protein phosphatase 1— targeted in many directions. J. Cell Sci. 115, 241–256 (2002).

    CAS  PubMed  Google Scholar 

  154. 154

    Kim, E., Goren, A. & Ast, G. Alternative splicing: current perspectives. Bioessays 30, 38–47 (2008).

    CAS  Google Scholar 

  155. 155

    Hallikas, O. et al. Genome-wide prediction of mammalian enhancers based on analysis of transcription-factor binding affinity. Cell 124, 47–59 (2006).

    CAS  PubMed  Google Scholar 

  156. 156

    Babu, M. M., Luscombe, N. M., Aravind, L., Gerstein, M. & Teichmann, S. A. Structure and evolution of transcriptional regulatory networks. Curr. Opin. Struct. Biol. 14, 283–291 (2004).

    CAS  Google Scholar 

  157. 157

    Tacke, R. & Manley, J. L. The human splicing factors ASF/SF2 and SC35 possess distinct, functionally significant RNA binding specificities. EMBO J. 14, 3540–3551 (1995).

    CAS  PubMed  PubMed Central  Google Scholar 

  158. 158

    Kent, O. A., Ritchie, D. B. & Macmillan, A. M. Characterization of a U2AF-independent commitment complex (E′) in the mammalian spliceosome assembly pathway. Mol. Cell. Biol. 25, 233–40 (2005).

    CAS  PubMed  PubMed Central  Google Scholar 

  159. 159

    Shen, H. & Green, M. R. RS domains contact splicing signals and promote splicing by a common mechanism in yeast through humans. Genes Dev. 20, 1755–1765 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  160. 160

    Huang, Y., Gattoni, R., Stevenin, J. & Steitz, J. A. SR splicing factors serve as adapter proteins for TAP-dependent mRNA export. Mol. Cell 11, 837–843 (2003).

    CAS  Google Scholar 

  161. 161

    Zhang, Z. & Krainer, A. R. Involvement of SR proteins in mRNA surveillance. Mol. Cell 16, 597–607 (2004).

    CAS  Google Scholar 

  162. 162

    Sanford, J. R., Gray, N. K., Beckmann, K. & Caceres, J. F. A novel role for shuttling SR proteins in mRNA translation. Genes Dev. 18, 755–768 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

  163. 163

    Cartegni, L. & Krainer, A. R. Disruption of an SF2/ASF-dependent exonic splicing enhancer in SMN2 causes spinal muscular atrophy in the absence of SMN1. Nature Genet. 30, 377–384 (2002).

    CAS  PubMed  Google Scholar 

  164. 164

    Cartegni, L., Hastings, M. L., Calarco, J. A., de Stanchina, E. & Krainer, A. R. Determinants of exon 7 splicing in the spinal muscular atrophy genes, SMN1 and SMN2. Am. J. Hum. Genet. 78, 63–77 (2006).

    CAS  Google Scholar 

Download references


Work from the authors' laboratory was supported in part by grants from the National Institutes of Health. We thank C. David for comments on the manuscript.

Author information



Corresponding author

Correspondence to James L. Manley.

Related links

Related links


James L. Manley's homepage


Small nuclear ribonucleoprotein particle

(snRNP). A protein, including U1, U2, U4, U5 and U6, which contains U-rich small nuclear RNAs (snRNAs) and both small nuclear ribonucleoprotein (snRNP)-specific and common proteins, and is a core component of the spliceosome.

Branch point

A nucleotide, usually an adenosine, within a variably conserved branch point sequence upstream of the 3′ splice site, the 2′ hydroxyl group of which attacks the 5′ splice site in the first step of splicing.

SR (Ser–Arg) protein family

A family of nuclear factors that have many important roles in splicing mRNA precursors in metazoan organisms, functioning in both constitutive and alternative splicing.

Heterologous nuclear RNP

(hnRNP). A pre-mRNA- or mRNA-binding protein that associates with transcripts during or after transcription and influences their function and fate. Some hnRNPs shuttle in and out of nuclei, whereas others are constitutively nuclear.

Alternative exon

An exon that is included in mature mRNA in certain cellular contexts but excluded in others.

RS (Arg–Ser repeat-containing) domain

A protein domain that is variable in length and enriched in Arg–Ser dipeptides and seems to be involved in protein–protein and protein–RNA interactions.

Hu/ELAV family protein

A protein belonging to a family of nervous system-specific RNA-binding proteins that specifically bind to AU-rich sequences.


A method that combines cross-linking and immunoprecipitation to identify in vivo targets of RNA-binding proteins.

RRM domain

(RNA recognition motif domain). A protein domain that is frequently involved in sequence-specific single-stranded RNA binding. Also known as an RNP-type RNA-binding domain.

14-3-3 protein

A protein belonging to a family of conserved proteins that bind to phosphorylated serine and threonine residues and that are encoded by seven genes in most mammals. They bind diverse regulatory proteins, including kinases, phosphatases and transmembrane receptors.


A technique to determine the DNA or RNA sequence that is specifically recognized by a protein. The method involves multiple rounds of binding to an initially random sequence until a high-affinity consensus sequence emerges.

Rights and permissions

Reprints and Permissions

About this article

Cite this article

Chen, M., Manley, J. Mechanisms of alternative splicing regulation: insights from molecular and genomics approaches. Nat Rev Mol Cell Biol 10, 741–754 (2009).

Download citation

Further reading


Quick links

Nature Briefing

Sign up for the Nature Briefing newsletter — what matters in science, free to your inbox daily.

Get the most important science stories of the day, free in your inbox. Sign up for Nature Briefing