Review Article | Published:

Eukaryotic core promoters and the functional basis of transcription initiation

Nature Reviews Molecular Cell Biologyvolume 19pages621637 (2018) | Download Citation

Abstract

RNA polymerase II (Pol II) core promoters are specialized DNA sequences at transcription start sites of protein-coding and non-coding genes that support the assembly of the transcription machinery and transcription initiation. They enable the highly regulated transcription of genes by selectively integrating regulatory cues from distal enhancers and their associated regulatory proteins. In this Review, we discuss the defining properties of gene core promoters, including their sequence features, chromatin architecture and transcription initiation patterns. We provide an overview of molecular mechanisms underlying the function and regulation of core promoters and their emerging functional diversity, which defines distinct transcription programmes. On the basis of the established properties of gene core promoters, we discuss transcription start sites within enhancers and integrate recent results obtained from dedicated functional assays to propose a functional model of transcription initiation. This model can explain the nature and function of transcription initiation at gene starts and at enhancers and can explain the different roles of core promoters, of Pol II and its associated factors and of the activating cues provided by enhancers and the transcription factors and cofactors they recruit.

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References

  1. 1.

    Spitz, F. & Furlong, E. E. M. Transcription factors: from enhancer binding to developmental control. Nat. Rev. Genet. 13, 613–626 (2012).

  2. 2.

    Levine, M. & Tjian, R. Transcription regulation and animal diversity. Nature 424, 147–151 (2003).

  3. 3.

    Levine, M., Cattoglio, C. & Tjian, R. Looping back to leap forward: transcription enters a new era. Cell 157, 13–25 (2014).

  4. 4.

    Herz, H.-M., Hu, D. & Shilatifard, A. Enhancer malfunction in cancer. Mol. Cell 53, 859–866 (2014).

  5. 5.

    Hampsey, M. Molecular genetics of the RNA polymerase II general transcriptional machinery. Microbiol. Mol. Biol. Rev. 62, 465–503 (1998).

  6. 6.

    Kadonaga, J. T. Perspectives on the RNA polymerase II core promoter. Wiley Interdiscip. Rev. Dev. Biol. 1, 40–51 (2012).

  7. 7.

    Banerji, J., Rusconi, S. & Schaffner, W. Expression of a beta-globin gene is enhanced by remote SV40 DNA sequences. Cell 27, 299–308 (1981).

  8. 8.

    Shlyueva, D., Stampfel, G. & Stark, A. Transcriptional enhancers: from properties to genome-wide predictions. Nat. Rev. Genet. 15, 272–286 (2014).

  9. 9.

    Zabidi, M. A. & Stark, A. Regulatory enhancer–core- promoter communication via transcription factors and cofactors. Trends Genet. 32, 801–814 (2016).

  10. 10.

    Shiraki, T. et al. Cap analysis gene expression for high-throughput analysis of transcriptional starting point and identification of promoter usage. Proc. Natl Acad. Sci. USA 100, 15776–15781 (2003).

  11. 11.

    Gu, W. et al. CapSeq and CIP-TAP identify Pol II start sites and reveal capped small RNAs as C. elegans piRNA precursors. Cell 151, 1488–1500 (2012).

  12. 12.

    Ni, T. et al. A paired-end sequencing strategy to map the complex landscape of transcription initiation. Nat. Methods 7, 521–527 (2010).

  13. 13.

    Nechaev, S. et al. Global analysis of short RNAs reveals widespread promoter-proximal stalling and arrest of Pol II in Drosophila. Science 327, 335–338 (2010).

  14. 14.

    Lam, M. T. Y. et al. Rev-Erbs repress macrophage gene expression by inhibiting enhancer-directed transcription. Nature 498, 511–515 (2013).

  15. 15.

    Core, L. J. et al. Analysis of nascent RNA identifies a unified architecture of initiation regions at mammalian promoters and enhancers. Nat. Genet. 46, 1311–1320 (2014). This work proposes a unified model of transcription initiation at promoters and enhancers and emphasizes that post-initiation transcript stability is the main distinction between the two elements.

  16. 16.

    Kwak, H., Fuda, N. J., Core, L. J. & Lis, J. T. Precise maps of RNA polymerase reveal how promoters direct initiation and pausing. Science 339, 950–953 (2013).

  17. 17.

    Affymetrix/Cold Spring Harbor Laboratory ENCODE Transcriptome Project. Post-transcriptional processing generates a diversity of 5ʹ- modified long and short RNAs. Nature 457, 1028–1032 (2009).

  18. 18.

    The FANTOM Consortium and RIKEN Genome Exploration Research Group and Genome Science Group (Genome Network Project Core Group). The transcriptional landscape of the mammalian genome. Science 309, 1559–1563 (2005).

  19. 19.

    Hoskins, R. A. et al. Genome-wide analysis of promoter architecture in Drosophila melanogaster. Genome Res. 21, 182–192 (2011).

  20. 20.

    Chen, R. A.-J. et al. The landscape of RNA polymerase II transcription initiation in C. elegans reveals promoter and enhancer architectures. Genome Res. 23, 1339–1347 (2013).

  21. 21.

    Haberle, V. et al. Two independent transcription initiation codes overlap on vertebrate core promoters. Nature 507, 381–385 (2014). This study reveals a widespread switch in TSS usage associated with distinct sequence properties during early embryonic development of zebrafish.

  22. 22.

    The FANTOM Consortium, The RIKEN PMI & CLST (DGT). A promoter-level mammalian expression atlas. Nature 507, 462–470 (2014).

  23. 23.

    The ENCODE Project Consortium et al. Identification and analysis of functional elements in 1% of the human genome by the ENCODE pilot project. Nature 447, 799–816 (2007).

  24. 24.

    Kapranov, P. et al. RNA maps reveal new RNA classes and a possible function for pervasive transcription. Science 316, 1484–1488 (2007).

  25. 25.

    Kim, T.-K. et al. Widespread transcription at neuronal activity-regulated enhancers. Nature 465, 182–187 (2010). This study reports widespread bidirectional transcription from enhancers giving rise to eRNAs.

  26. 26.

    Andersson, R. et al. An atlas of active enhancers across human cell types and tissues. Nature 507, 455–461 (2014). This study uses bidirectional transcription initiation to predict enhancers and their activity across numerous human cell types.

  27. 27.

    De Santa, F. et al. A large fraction of extragenic RNA pol II transcription sites overlap enhancers. PLoS Biol. 8, e1000384 (2010).

  28. 28.

    Koch, F. et al. Transcription initiation platforms and GTF recruitment at tissue-specific enhancers and promoters. Nat. Struct. Mol. Biol. 18, 956–963 (2011).

  29. 29.

    Arner, E. et al. Transcribed enhancers lead waves of coordinated transcription in transitioning mammalian cells. Science 347, 1010–1014 (2015).

  30. 30.

    Li, W. et al. Functional roles of enhancer RNAs for oestrogen-dependent transcriptional activation. Nature 498, 516–520 (2013).

  31. 31.

    Schaukowitch, K. et al. Enhancer RNA facilitates NELF release from immediate early genes. Mol. Cell 56, 29–42 (2014).

  32. 32.

    Andersson, R. et al. Nuclear stability and transcriptional directionality separate functionally distinct RNA species. Nat. Commun. 5, 5336 (2014).

  33. 33.

    Seila, A. C. et al. Divergent transcription from active promoters. Science 322, 1849–1851 (2008).

  34. 34.

    Preker, P. et al. RNA exosome depletion reveals transcription upstream of active human promoters. Science 322, 1851–1854 (2008).

  35. 35.

    Core, L. J., Waterfall, J. J. & Lis, J. T. Nascent RNA sequencing reveals widespread pausing and divergent initiation at human promoters. Science 322, 1845–1848 (2008). References 33–35 report widespread antisense transcription from gene promoters giving rise to short unstable upstream antisense RNAs.

  36. 36.

    Andersson, R. et al. Human gene promoters are intrinsically bidirectional. Mol. Cell 60, 346–347 (2015).

  37. 37.

    Duttke, S. H. C. et al. Human promoters are intrinsically directional. Mol. Cell 57, 674–684 (2015).

  38. 38.

    Scruggs, B. S. et al. Bidirectional transcription arises from two distinct hubs of transcription factor binding and active chromatin. Mol. Cell 58, 1101–1112 (2015). References 37 and 38 demonstrate that bidirectional transcription from promoters arises from two separate and intrinsically directional transcription complexes.

  39. 39.

    Sigova, A. A. et al. Divergent transcription of long noncoding RNA/mRNA gene pairs in embryonic stem cells. Proc. Natl Acad. Sci. USA 110, 2876–2881 (2013).

  40. 40.

    Lepoivre, C. et al. Divergent transcription is associated with promoters of transcriptional regulators. BMC Genomics 14, 914 (2013).

  41. 41.

    Andersson, R. Promoter or enhancer, what’s the difference? Deconstruction of established distinctions and presentation of a unifying model. Bioessays 37, 314–323 (2015).

  42. 42.

    Kim, T.-K. & Shiekhattar, R. Architectural and functional commonalities between enhancers and promoters. Cell 162, 948–959 (2015).

  43. 43.

    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).

  44. 44.

    Ntini, E. et al. Polyadenylation site-induced decay of upstream transcripts enforces promoter directionality. Nat. Struct. Mol. Biol. 20, 923–928 (2013).

  45. 45.

    Carninci, P. et al. Genome-wide analysis of mammalian promoter architecture and evolution. Nat. Genet. 38, 626–635 (2006). This work uses genome-wide maps of human and mouse TSSs to describe two classes of promoters that differ in initiation pattern: focused TATA-box-enriched promoters and broad CpG-rich promoters.

  46. 46.

    Lenhard, B., Sandelin, A. & Carninci, P. Metazoan promoters: emerging characteristics and insights into transcriptional regulation. Nat. Rev. Genet. 13, 233–245 (2012).

  47. 47.

    Schor, I. E. et al. Promoter shape varies across populations and affects promoter evolution and expression noise. Nat. Genet. 49, 550–558 (2017). This work identifies natural genetic variants that affect TSS distributions within core promoters in flies.

  48. 48.

    Lifton, R. P., Goldberg, M. L., Karp, R. W. & Hogness, D. S. The organization of the histone genes in Drosophila melanogaster: functional and evolutionary implications. Cold Spring Harb. Symp. Quant. Biol. 42, 1047–1051 (1978).

  49. 49.

    Goldberg, M. L. Sequence Analysis of Drosophila Histone Genes. Thesis, Stanford Univ. (1979).

  50. 50.

    Ponjavic, J. et al. Transcriptional and structural impact of TATA-initiation site spacing in mammalian core promoters. Genome Biol. 7, R78 (2006).

  51. 51.

    Ohler, U., Liao, G.-C., Niemann, H. & Rubin, G. M. Computational analysis of core promoters in the Drosophila genome. Genome Biol. 3, RESEARCH0087 (2002).

  52. 52.

    FitzGerald, P. C., Sturgill, D., Shyakhtenko, A., Oliver, B. & Vinson, C. Comparative genomics of drosophila and human core promoters. Genome Biol. 7, R53 (2006). References 51 and 52 provide comprehensive computational analyses of over-represented sequence motifs in fly and human core promoters.

  53. 53.

    Patikoglou, G. A. et al. TATA element recognition by the TATA box-binding protein has been conserved throughout evolution. Genes Dev. 13, 3217–3230 (1999).

  54. 54.

    Burley, S. K. & Roeder, R. G. Biochemistry and structural biology of transcription factor IID (TFIID). Annu. Rev. Biochem. 65, 769–799 (1996).

  55. 55.

    Louder, R. K. et al. Structure of promoter-bound TFIID and model of human pre-initiation complex assembly. Nature 531, 604–609 (2016). This work reports a structure of the human promoter-bound TFIID, revealing contacts between TFIID subunits and specific core-promoter motifs.

  56. 56.

    Smale, S. T. & Baltimore, D. The ‘initiator’ as a transcription control element. Cell 57, 103–113 (1989).

  57. 57.

    Chalkley, G. E. & Verrijzer, C. P. DNA binding site selection by RNA polymerase II TAFs: a TAF(II)250-TAF(II)150 complex recognizes the initiator. EMBO J 18, 4835–4845 (1999).

  58. 58.

    Vo Ngoc, L., Cassidy, C. J., Huang, C. Y., Duttke, S. H. C. & Kadonaga, J. T. The human initiator is a distinct and abundant element that is precisely positioned in focused core promoters. Genes Dev. 31, 6–11 (2017).

  59. 59.

    Burke, T. W. & Kadonaga, J. T. Drosophila TFIID binds to a conserved downstream basal promoter element that is present in many TATA-box-deficient promoters. Genes Dev. 10, 711–724 (1996).

  60. 60.

    Burke, T. W. & Kadonaga, J. T. The downstream core promoter element, DPE, is conserved from Drosophila to humans and is recognized by TAFII60 of Drosophila. Genes Dev. 11, 3020–3031 (1997).

  61. 61.

    Kutach, A. K. & Kadonaga, J. T. The downstream promoter element DPE appears to be as widely used as the TATA box in Drosophila core promoters. Mol. Cell. Biol. 20, 4754–4764 (2000).

  62. 62.

    Engstrom, P. G., Ho Sui, S. J., Drivenes, O., Becker, T. S. & Lenhard, B. Genomic regulatory blocks underlie extensive microsynteny conservation in insects. Genome Res. 17, 1898–1908 (2007).

  63. 63.

    Lim, C. Y. et al. The MTE, a new core promoter element for transcription by RNA polymerase II. Genes Dev. 18, 1606–1617 (2004).

  64. 64.

    Lagrange, T., Kapanidis, A. N., Tang, H., Reinberg, D. & Ebright, R. H. New core promoter element in RNA polymerase II-dependent transcription: sequence-specific DNA binding by transcription factor IIB. Genes Dev. 12, 34–44 (1998).

  65. 65.

    Deng, W. & Roberts, S. G. E. A core promoter element downstream of the TATA box that is recognized by TFIIB. Genes Dev. 19, 2418–2423 (2005).

  66. 66.

    Lewis, B. A., Kim, T. K. & Orkin, S. H. A downstream element in the human beta-globin promoter: evidence of extended sequence-specific transcription factor IID contacts. Proc. Natl Acad. Sci. USA 97, 7172–7177 (2000).

  67. 67.

    Lee, D.-H. et al. Functional characterization of core promoter elements: the downstream core element is recognized by TAF1. Mol. Cell. Biol. 25, 9674–9686 (2005).

  68. 68.

    Rach, E. A., Yuan, H.-Y., Majoros, W. H., Tomancak, P. & Ohler, U. Motif composition, conservation and condition-specificity of single and alternative transcription start sites in the drosophila genome. Genome Biol. 10, R73 (2009). This work shows differential motif enrichment and spatiotemporal utilization of core promoters with focused and dispersed initiation patterns in flies.

  69. 69.

    Mikhaylichenko, O. et al. The degree of enhancer or promoter activity is reflected by the levels and directionality of eRNA transcription. Genes Dev. 32, 42–57 (2018). This work introduces a functional assay to simultaneously measure enhancer and promoter activity of candidate fragments and shows that core-promoter motifs confer promoter functionality to enhancers.

  70. 70.

    Juven-Gershon, T., Cheng, S. & Kadonaga, J. T. Rational design of a super core promoter that enhances gene expression. Nat. Methods 3, 917–922 (2006).

  71. 71.

    Pfeiffer, B. D. et al. Tools for neuroanatomy and neurogenetics in Drosophila. Proc. Natl Acad. Sci. USA 105, 9715–9720 (2008).

  72. 72.

    Even, D. Y. et al. Engineered promoters for potent transient overexpression. PLoS ONE 11, e0148918 (2016).

  73. 73.

    Gardiner-Garden, M. & Frommer, M. CpG islands in vertebrate genomes. J. Mol. Biol. 196, 261–282 (1987).

  74. 74.

    Saxonov, S., Berg, P. & Brutlag, D. L. A genome-wide analysis of CpG dinucleotides in the human genome distinguishes two distinct classes of promoters. Proc. Natl Acad. Sci. USA 103, 1412–1417 (2006).

  75. 75.

    Akalin, A. et al. Transcriptional features of genomic regulatory blocks. Genome Biol. 10, R38 (2009).

  76. 76.

    Dreos, R., Ambrosini, G. & Bucher, P. Influence of rotational nucleosome positioning on transcription start site selection in animal promoters. PLoS Comput. Biol. 12, e1005144 (2016).

  77. 77.

    Satchwell, S. C., Drew, H. R. & Travers, A. A. Sequence periodicities in chicken nucleosome core DNA. J. Mol. Biol. 191, 659–675 (1986).

  78. 78.

    Widom, J. Role of DNA sequence in nucleosome stability and dynamics. Q. Rev. Biophys. 34, 269–324 (2001).

  79. 79.

    Segal, E. et al. A genomic code for nucleosome positioning. Nature 442, 772–778 (2006).

  80. 80.

    Yuan, G.-C. et al. Genome-scale identification of nucleosome positions in S. cerevisiae. Science 309, 626–630 (2005).

  81. 81.

    Mavrich, T. N. et al. Nucleosome organization in the Drosophila genome. Nature 453, 358–362 (2008). This work maps nucleosome positions across a metazoan genome and reveals the organization of nucleosomes around active promoters.

  82. 82.

    Jiang, C. & Pugh, B. F. Nucleosome positioning and gene regulation: advances through genomics. Nat. Rev. Genet. 10, 161–172 (2009).

  83. 83.

    Jin, C. et al. H3.3/H2A. Z double variant-containing nucleosomes mark ‘nucleosome-free regions’ of active promoters and other regulatory regions. Nat. Genet. 41, 941–945 (2009).

  84. 84.

    Fei, J. et al. The prenucleosome, a stable conformational isomer of the nucleosome. Genes Dev. 29, 2563–2575 (2015).

  85. 85.

    Henikoff, J. G., Belsky, J. A., Krassovsky, K., MacAlpine, D. M. & Henikoff, S. Epigenome characterization at single base-pair resolution. Proc. Natl Acad. Sci. USA 108, 18318–18323 (2011).

  86. 86.

    Rhee, H. S., Bataille, A. R., Zhang, L. & Pugh, B. F. Subnucleosomal structures and nucleosome asymmetry across a genome. Cell 159, 1377–1388 (2014).

  87. 87.

    Mueller, B. et al. Widespread changes in nucleosome accessibility without changes in nucleosome occupancy during a rapid transcriptional induction. Genes Dev. 31, 451–462 (2017).

  88. 88.

    Mieczkowski, J. et al. MNase titration reveals differences between nucleosome occupancy and chromatin accessibility. Nat. Commun. 7, 11485 (2016).

  89. 89.

    Rach, E. A. et al. Transcription initiation patterns indicate divergent strategies for gene regulation at the chromatin level. PLoS Genet. 7, e1001274 (2011).

  90. 90.

    Kubik, S. et al. Nucleosome stability distinguishes two different promoter types at all protein-coding genes in yeast. Mol. Cell 60, 422–434 (2015).

  91. 91.

    Zaret, K. S. & Carroll, J. S. Pioneer transcription factors: establishing competence for gene expression. Genes Dev. 25, 2227–2241 (2011).

  92. 92.

    Shimojima, T. et al. Drosophila FACT contributes to Hox gene expression through physical and functional interactions with GAGA factor. Genes Dev. 17, 1605–1616 (2003).

  93. 93.

    Fuda, N. J. et al. GAGA factor maintains nucleosome-free regions and has a role in RNA polymerase II recruitment to promoters. PLoS Genet. 11, e1005108 (2015).

  94. 94.

    Weber, C. M., Ramachandran, S. & Henikoff, S. Nucleosomes are context-specific, H2A. Z-modulated barriers to RNA polymerase. Mol. Cell 53, 819–830 (2014).

  95. 95.

    Mousavi, K. et al. eRNAs promote transcription by establishing chromatin accessibility at defined genomic loci. Mol. Cell 51, 606–617 (2013).

  96. 96.

    Gilchrist, D. A. et al. Pausing of RNA polymerase II disrupts DNA-specified nucleosome organization to enable precise gene regulation. Cell 143, 540–551 (2010).

  97. 97.

    Ahmad, K. & Henikoff, S. The histone variant H3.3 marks active chromatin by replication-independent nucleosome assembly. Mol. Cell 9, 1191–1200 (2002).

  98. 98.

    Pradhan, S. K. et al. EP400 deposits H3.3 into promoters and enhancers during gene activation. Mol. Cell 61, 27–38 (2016).

  99. 99.

    Raisner, R. M. et al. Histone variant H2A. Z marks the 5ʹ ends of both active and inactive genes in euchromatin. Cell 123, 233–248 (2005).

  100. 100.

    Barski, A. et al. High-resolution profiling of histone methylations in the human genome. Cell 129, 823–837 (2007). This study maps different histone modifications genome-wide and identified those associated with active or repressed promoters.

  101. 101.

    Ng, H.-H., Robert, F., Young, R. A. & Struhl, K. Targeted recruitment of Set1 histone methylase by elongating Pol II provides a localized mark and memory of recent transcriptional activity. Mol. Cell 11, 709–719 (2003).

  102. 102.

    Hathaway, N. A. et al. Dynamics and memory of heterochromatin in living cells. Cell 149, 1447–1460 (2012).

  103. 103.

    Zhao, R., Nakamura, T., Fu, Y., Lazar, Z. & Spector, D. L. Gene bookmarking accelerates the kinetics of post-mitotic transcriptional re-activation. Nat. Cell Biol. 13, 1295–1304 (2011).

  104. 104.

    Tropberger, P. et al. Regulation of transcription through acetylation of H3K122 on the lateral surface of the histone octamer. Cell 152, 859–872 (2013).

  105. 105.

    Neumann, H. et al. A method for genetically installing site-specific acetylation in recombinant histones defines the effects of H3 K56 acetylation. Mol. Cell 36, 153–163 (2009).

  106. 106.

    Tessarz, P. & Kouzarides, T. Histone core modifications regulating nucleosome structure and dynamics. Nat. Rev. Mol. Cell Biol. 15, 703–708 (2014).

  107. 107.

    Dey, A., Chitsaz, F., Abbasi, A., Misteli, T. & Ozato, K. The double bromodomain protein Brd4 binds to acetylated chromatin during interphase and mitosis. Proc. Natl Acad. Sci. USA 100, 8758–8763 (2003).

  108. 108.

    Hödl, M. & Basler, K. Transcription in the absence of histone H3.2 and H3K4 methylation. Curr. Biol. 22, 2253–2257 (2012).

  109. 109.

    Hödl, M. & Basler, K. Transcription in the absence of histone H3.3. Curr. Biol. 19, 1221–1226 (2009).

  110. 110.

    Pengelly, A. R., Copur, Ö., Jäckle, H., Herzig, A. & Müller, J. A histone mutant reproduces the phenotype caused by loss of histone-modifying factor Polycomb. Science 339, 698–699 (2013).

  111. 111.

    Sterner, D. E. & Berger, S. L. Acetylation of histones and transcription-related factors. Microbiol. Mol. Biol. Rev. 64, 435–459 (2000).

  112. 112.

    Imhof, A. et al. Acetylation of general transcription factors by histone acetyltransferases. Curr. Biol. 7, 689–692 (1997).

  113. 113.

    Roe, J.-S., Mercan, F., Rivera, K., Pappin, D. J. & Vakoc, C. R. BET bromodomain inhibition suppresses the function of hematopoietic transcription factors in acute myeloid leukemia. Mol. Cell 58, 1028–1039 (2015).

  114. 114.

    Schröder, S. et al. Acetylation of RNA polymerase II regulates growth-factor-induced gene transcription in mammalian cells. Mol. Cell 52, 314–324 (2013).

  115. 115.

    Rickels, R. et al. Histone H3K4 monomethylation catalyzed by Trr and mammalian COMPASS-like proteins at enhancers is dispensable for development and viability. Nat. Genet. 156, 645–1653 (2017).

  116. 116.

    Dorighi, K. M. et al. Mll3 and Mll4 facilitate enhancer RNA synthesis and transcription from promoters independently of H3K4 monomethylation. Mol. Cell 66, 568–576.e4 (2017).

  117. 117.

    Pollex, T. & Furlong, E. E. M. Correlation does not imply causation: histone methyltransferases, but not histone methylation, SET the stage for enhancer activation. Mol. Cell 66, 439–441 (2017).

  118. 118.

    Andersen, P. R., Tirian, L., Vunjak, M. & Brennecke, J. A heterochromatin-dependent transcription machinery drives piRNA expression. Nature 549, 54–59 (2017). This study shows that histone modifications recruit the transcription machinery to transcribe heterochromatic loci that are a source of small RNAs.

  119. 119.

    Thomas, M. C. & Chiang, C.-M. The general transcription machinery and general cofactors. Crit. Rev. Biochem. Mol. Biol. 41, 105–178 (2006).

  120. 120.

    Orphanides, G., Lagrange, T. & Reinberg, D. The general transcription factors of RNA polymerase II. Genes Dev. 10, 2657–2683 (1996).

  121. 121.

    Sainsbury, S., Bernecky, C. & Cramer, P. Structural basis of transcription initiation by RNA polymerase II. Nat. Rev. Mol. Cell. Biol. 16, 129–143 (2015).

  122. 122.

    Zhang, Z. et al. Rapid dynamics of general transcription factor TFIIB binding during preinitiation complex assembly revealed by single-molecule analysis. Genes Dev. 30, 2106–2118 (2016).

  123. 123.

    He, Y. et al. Near-atomic resolution visualization of human transcription promoter opening. Nature 533, 359–365 (2016).

  124. 124.

    Plaschka, C. et al. Transcription initiation complex structures elucidate DNA opening. Nature 533, 353–358 (2016). This study reports structures of open and closed yeast PIC complexes and proposes a mechanism of DNA duplex opening.

  125. 125.

    Vermeulen, M. et al. Selective anchoring of TFIID to nucleosomes by trimethylation of histone H3 lysine 4. Cell 131, 58–69 (2007).

  126. 126.

    Papai, G. et al. TFIIA and the transactivator Rap1 cooperate to commit TFIID for transcription initiation. Nature 465, 956–960 (2010).

  127. 127.

    Liu, W.-L. et al. Structures of three distinct activator-TFIID complexes. Genes Dev. 23, 1510–1521 (2009).

  128. 128.

    Chopra, V. S. et al. Transcriptional activation by GAGA factor is through its direct interaction with dmTAF3. Dev. Biol. 317, 660–670 (2008).

  129. 129.

    Hochheimer, A. & Tjian, R. Diversified transcription initiation complexes expand promoter selectivity and tissue-specific gene expression. Genes Dev. 17, 1309–1320 (2003).

  130. 130.

    Taatjes, D. J., Marr, M. T. & Tjian, R. Regulatory diversity among metazoan co-activator complexes. Nat. Rev. Mol. Cell. Biol. 5, 403–410 (2004).

  131. 131.

    Goodrich, J. A. & Tjian, R. Unexpected roles for core promoter recognition factors in cell-type-specific transcription and gene regulation. Nat. Rev. Genet. 11, 549–558 (2010).

  132. 132.

    Jones, K. A. Changing the core of transcription. eLife 3, e03575 (2014).

  133. 133.

    Hochheimer, A., Zhou, S., Zheng, S., Holmes, M. C. & Tjian, R. TRF2 associates with DREF and directs promoter-selective gene expression in Drosophila. Nature 420, 439–445 (2002).

  134. 134.

    Wang, Y.-L. et al. TRF2, but not TBP, mediates the transcription of ribosomal protein genes. Genes Dev. 28, 1550–1555 (2014). References 133 and 134 reveal that TRF2 replaces TBP within the PIC to drive transcription of a specific subset of genes.

  135. 135.

    Isogai, Y., Keles, S., Prestel, M., Hochheimer, A. & Tjian, R. Transcription of histone gene cluster by differential core-promoter factors. Genes Dev. 21, 2936–2949 (2007).

  136. 136.

    Rhee, H. S. & Pugh, B. F. Genome-wide structure and organization of eukaryotic pre-initiation complexes. Nature 483, 295–301 (2012). This study maps positions of PIC components at high resolution across the entire yeast genome.

  137. 137.

    Basehoar, A. D., Zanton, S. J. & Pugh, B. F. Identification and distinct regulation of yeast TATA box-containing genes. Cell 116, 699–709 (2004).

  138. 138.

    Struhl, K. Constitutive and inducible Saccharomyces cerevisiae promoters: evidence for two distinct molecular mechanisms. Mol. Cell. Biol. 6, 3847–3853 (1986).

  139. 139.

    Baptista, T. et al. SAGA is a general cofactor for RNA polymerase II transcription. Mol. Cell 68, 1–20 (2017).

  140. 140.

    Warfield, L. et al. Transcription of nearly all yeast RNA polymerase II- transcribed genes is dependent on transcription factor TFIID. Mol. Cell 68, 1–18 (2017).

  141. 141.

    Zeitlinger, J. et al. RNA polymerase stalling at developmental control genes in the Drosophila melanogaster embryo. Nat. Genet. 39, 1512–1516 (2007).

  142. 142.

    Muse, G. W. et al. RNA polymerase is poised for activation across the genome. Nat. Genet. 39, 1507–1511 (2007). References 141 and 142 report widespread Pol II pausing at developmentally regulated genes in flies.

  143. 143.

    Guenther, M. G., Levine, S. S., Boyer, L. A., Jaenisch, R. & Young, R. A. A chromatin landmark and transcription initiation at most promoters in human cells. Cell 130, 77–88 (2007).

  144. 144.

    Rougvie, A. E. & Lis, J. T. The RNA polymerase II molecule at the 5ʹ end of the uninduced hsp70 gene of D. melanogaster is transcriptionally engaged. Cell 54, 795–804 (1988). This study shows that Pol II pauses downstream of the TSS.

  145. 145.

    Lis, J. T., Mason, P., Peng, J., Price, D. H. & Werner, J. P-TEFb kinase recruitment and function at heat shock loci. Genes Dev. 14, 792–803 (2000).

  146. 146.

    Henriques, T. et al. Stable pausing by RNA polymerase II provides an opportunity to target and integrate regulatory signals. Mol. Cell 52, 517–528 (2013).

  147. 147.

    Boettiger, A. N. & Levine, M. Synchronous and stochastic patterns of gene activation in the Drosophila embryo. Science 325, 471–473 (2009).

  148. 148.

    Lagha, M. et al. Paused Pol II coordinates tissue morphogenesis in the Drosophila embryo. Cell 153, 976–987 (2013).

  149. 149.

    Williams, L. H. et al. Pausing of RNA polymerase II regulates mammalian developmental potential through control of signaling networks. Mol. Cell 58, 311–322 (2015).

  150. 150.

    Jonkers, I., Kwak, H. & Lis, J. T. Genome-wide dynamics of Pol II elongation and its interplay with promoter proximal pausing, chromatin, and exons. eLife 3, e02407 (2014).

  151. 151.

    Shao, W. & Zeitlinger, J. Paused RNA polymerase II inhibits new transcriptional initiation. Nat. Genet. 16, 129–1051 (2017).

  152. 152.

    Krebs, A. R. et al. Genome-wide single-molecule footprinting reveals high RNA polymerase II turnover at paused promoters. Mol. Cell 67, 411–422.e4 (2017).

  153. 153.

    Gressel, S. et al. CDK9-dependent RNA polymerase II pausing controls transcription initiation. eLife 6, R106 (2017). References 151–153 report a wide-range of paused Pol II half-lives at promoters genome-wide.

  154. 154.

    Ehrensberger, A. H., Kelly, G. P. & Svejstrup, J. Q. Mechanistic interpretation of promoter-proximal peaks and RNAPII density maps. Cell 154, 713–715 (2013).

  155. 155.

    Hendrix, D. A., Hong, J.-W., Zeitlinger, J., Rokhsar, D. S. & Levine, M. S. Promoter elements associated with RNA Pol II stalling in the Drosophila embryo. Proc. Natl Acad. Sci. USA 105, 7762–7767 (2008).

  156. 156.

    Veloso, A. et al. Rate of elongation by RNA polymerase II is associated with specific gene features and epigenetic modifications. Genome Res. 24, 896–905 (2014).

  157. 157.

    Gartenberg, M. R. & Wang, J. C. Positive supercoiling of DNA greatly diminishes mRNA synthesis in yeast. Proc. Natl Acad. Sci. USA 89, 11461–11465 (1992).

  158. 158.

    Joshi, R. S., Piña, B. & Roca, J. Positional dependence of transcriptional inhibition by DNA torsional stress in yeast chromosomes. EMBO J. 29, 740–748 (2010).

  159. 159.

    Henriques, T. et al. Widespread transcriptional pausing and elongation control at enhancers. Genes Dev. 32, 26–41 (2018).

  160. 160.

    Chen, F. X. et al. PAF1 regulation of promoter-proximal pause release via enhancer activation. Science 357, 1294–1298 (2017). References 159 and 160 report widespread pausing of Pol II at enhancers.

  161. 161.

    Baranello, L. et al. RNA polymerase II regulates topoisomerase 1 activity to favor efficient transcription. Cell 165, 357–371 (2016).

  162. 162.

    Missra, A. & Gilmour, D. S. Interactions between DSIF (DRB sensitivity inducing factor), NELF (negative elongation factor), and the Drosophila RNA polymerase II transcription elongation complex. Proc. Natl Acad. Sci. USA 107, 11301–11306 (2010).

  163. 163.

    Yamaguchi, Y., Shibata, H. & Handa, H. Transcription elongation factors DSIF and NELF: promoter-proximal pausing and beyond. Biochim. Biophys. Acta 1829, 98–104 (2013).

  164. 164.

    Bernecky, C., Plitzko, J. M. & Cramer, P. Structure of a transcribing RNA polymerase II-DSIF complex reveals a multidentate DNA-RNA clamp. Nat. Struct. Mol. Biol. 24, 809–815 (2017).

  165. 165.

    Qiu, Y. & Gilmour, D. S. Identification of regions in the Spt5 subunit of DRB sensitivity-inducing factor (DSIF) that are involved in promoter-proximal pausing. J. Biol. Chem. 292, 5555–5570 (2017). References 164 and 165 provide structural and biochemical evidence that DSIF contacts nascent RNA protruding from Pol II, ascribing it a role in triggering Pol II pausing.

  166. 166.

    Ehara, H. et al. Structure of the complete elongation complex of RNA polymerase II with basal factors. Science 357, 921–924 (2017).

  167. 167.

    Li, G. et al. Extensive promoter-centered chromatin interactions provide a topological basis for transcription regulation. Cell 148, 84–98 (2012).

  168. 168.

    Rao, S. S. P. et al. A 3D map of the human genome at kilobase resolution reveals principles of chromatin looping. Cell 159, 1665–1680 (2014).

  169. 169.

    Sanyal, A., Lajoie, B. R., Jain, G. & Dekker, J. The long-range interaction landscape of gene promoters. Nature 489, 109–113 (2012).

  170. 170.

    Beagrie, R. A. et al. Complex multi-enhancer contacts captured by genome architecture mapping. Nature 543, 519–524 (2017).

  171. 171.

    Symmons, O. et al. Functional and topological characteristics of mammalian regulatory domains. Genome Res. 24, 390–400 (2014).

  172. 172.

    Merkenschlager, M. & Nora, E. P. CTCF and cohesin in genome folding and transcriptional gene regulation. Annu. Rev. Genomics Hum. Genet. 17, 17–43 (2016).

  173. 173.

    Spitz, F. Gene regulation at a distance: from remote enhancers to 3D regulatory ensembles. Semin. Cell Dev. Biol. 57, 57–67 (2016).

  174. 174.

    Ghavi-Helm, Y. et al. Enhancer loops appear stable during development and are associated with paused polymerase. Nature 512, 96–100 (2014).

  175. 175.

    Michel, M. & Cramer, P. Transitions for regulating early transcription. Cell 153, 943–944 (2013).

  176. 176.

    Eychenne, T. et al. Functional interplay between mediator and TFIIB in preinitiation complex assembly in relation to promoter architecture. Genes Dev. 30, 2119–2132 (2016).

  177. 177.

    Esnault, C. et al. Mediator-dependent recruitment of TFIIH modules in preinitiation complex. Mol. Cell 31, 337–346 (2008).

  178. 178.

    Visel, A. et al. ChIP-seq accurately predicts tissue-specific activity of enhancers. Nature 457, 854–858 (2009).

  179. 179.

    Boija, A. et al. CBP regulates recruitment and release of promoter-proximal RNA polymerase II. Mol. Cell 68, 491–503.e5 (2017).

  180. 180.

    Sawado, T., Halow, J., Bender, M. A. & Groudine, M. The beta -globin locus control region (LCR) functions primarily by enhancing the transition from transcription initiation to elongation. Genes Dev. 17, 1009–1018 (2003).

  181. 181.

    Yang, Z. et al. Recruitment of P-TEFb for stimulation of transcriptional elongation by the bromodomain protein Brd4. Mol. Cell 19, 535–545 (2005).

  182. 182.

    Jang, M. K. et al. The bromodomain protein Brd4 is a positive regulatory component of P-TEFb and stimulates RNA polymerase II-dependent transcription. Mol. Cell 19, 523–534 (2005).

  183. 183.

    Zuber, J. et al. RNAi screen identifies Brd4 as a therapeutic target in acute myeloid leukaemia. Nature 478, 524–528 (2011).

  184. 184.

    Delmore, J. E. et al. BET bromodomain inhibition as a therapeutic strategy to target c-Myc. Cell 146, 904–917 (2011). References 183 and 184 identify BRD4 as a gene-specific regulator whose depletion affects only a subset of genes.

  185. 185.

    Lovén, J. et al. Selective inhibition of tumor oncogenes by disruption of super-enhancers. Cell 153, 320–334 (2013).

  186. 186.

    Rathert, P. et al. Transcriptional plasticity promotes primary and acquired resistance to BET inhibition. Nature 525, 543–547 (2015).

  187. 187.

    Winter, G. E. et al. BET bromodomain proteins function as master transcription elongation factors independent of CDK9 recruitment. Mol. Cell 67, 5–18 (2017).

  188. 188.

    Muhar, M. et al. SLAM-seq defines direct gene-regulatory functions of the BRD4-MYC axis. Science https://doi.org/10.1126/science.aao2793 (2018). References 187 and 188 demonstrate that BRD proteins, in particular BRD4, are globally required for transition into productive elongation in a manner that is independent of CDK9 recruitment.

  189. 189.

    Chen, F. X. et al. PAF1, a molecular regulator of promoter-proximal pausing by RNA polymerase II. Cell 162, 1003–1015 (2015).

  190. 190.

    Chubb, J. R., Trcek, T., Shenoy, S. M. & Singer, R. H. Transcriptional pulsing of a developmental gene. Curr. Biol. 16, 1018–1025 (2006).

  191. 191.

    Raj, A., Peskin, C. S., Tranchina, D., Vargas, D. Y. & Tyagi, S. Stochastic mRNA synthesis in mammalian cells. PLoS Biol. 4, e309 (2006).

  192. 192.

    Tantale, K. et al. A single-molecule view of transcription reveals convoys of RNA polymerases and multi-scale bursting. Nat. Commun. 7, 12248 (2016).

  193. 193.

    Fukaya, T., Lim, B. & Levine, M. Enhancer control of transcriptional bursting. Cell 166, 358–368 (2016). This study provides evidence that enhancers regulate the frequency of transcription bursts synchronously from multiple promoters in their vicinity.

  194. 194.

    Bartman, C. R., Hsu, S. C., Hsiung, C. C.-S., Raj, A. & Blobel, G. A. Enhancer regulation of transcriptional bursting parameters revealed by forced chromatin looping. Mol. Cell 62, 237–247 (2016).

  195. 195.

    Hornung, G. et al. Noise-mean relationship in mutated promoters. Genome Res. 22, 2409–2417 (2012).

  196. 196.

    Blake, W. J. et al. Phenotypic consequences of promoter-mediated transcriptional noise. Mol. Cell 24, 853–865 (2006).

  197. 197.

    Tirosh, I., Weinberger, A., Carmi, M. & Barkai, N. A genetic signature of interspecies variations in gene expression. Nat. Genet. 38, 830–834 (2006).

  198. 198.

    Arnold, C. D. et al. Genome-wide assessment of sequence-intrinsic enhancer responsiveness at single-base-pair resolution. Nat. Biotechnol. 35, 136–144 (2016). This study measures enhancer responsiveness for all core promoters across the fly genome and demonstrates that core promoters show differential responses to different enhancers.

  199. 199.

    Deng, W. et al. Controlling long-range genomic interactions at a native locus by targeted tethering of a looping factor. Cell 149, 1233–1244 (2012).

  200. 200.

    Butler, J. E. & Kadonaga, J. T. Enhancer-promoter specificity mediated by DPE or TATA core promoter motifs. Genes Dev. 15, 2515–2519 (2001). This work demonstrates that TATA-box-containing and DPE-containing core promoters can be differentially activated when integrated into the same genomic locus.

  201. 201.

    Zabidi, M. A. et al. Enhancer-core-promoter specificity separates developmental and housekeeping gene regulation. Nature 518, 556–559 (2015). This work provides evidence for sequence-encoded enhancer–core-promoter specificity that distinguishes between housekeeping and developmental transcription programmes in the fly genome.

  202. 202.

    Ptashne, M. & Gann, A. Transcriptional activation by recruitment. Nature 386, 569–577 (1997).

  203. 203.

    Brent, R. & Ptashne, M. A eukaryotic transcriptional activator bearing the DNA specificity of a prokaryotic repressor. Cell 43, 729–736 (1985).

  204. 204.

    Hope, I. A. & Struhl, K. Functional dissection of a eukaryotic transcriptional activator protein, GCN4 of yeast. Cell 46, 885–894 (1986).

  205. 205.

    Keung, A. J., Bashor, C. J., Kiriakov, S., Collins, J. J. & Khalil, A. S. Using targeted chromatin regulators to engineer combinatorial and spatial transcriptional regulation. Cell 158, 110–120 (2014).

  206. 206.

    Stampfel, G. et al. Transcriptional regulators form diverse groups with context-dependent regulatory functions. Nature 528, 147–151 (2015). References 202–206 show that artificial recruitment of transcription factors and cofactors can be sufficient to drive transcription and that their activity is often context-dependent.

  207. 207.

    Juven-Gershon, T., Hsu, J.-Y. & Kadonaga, J. T. Caudal, a key developmental regulator, is a DPE-specific transcriptional factor. Genes Dev. 22, 2823–2830 (2008).

  208. 208.

    van Arensbergen, J., van Steensel, B. & Bussemaker, H. J. In search of the determinants of enhancer-promoter interaction specificity. Trends Cell Biol. 24, 695–702 (2014).

  209. 209.

    Petrenko, N., Jin, Y., Wong, K. H. & Struhl, K. Evidence that mediator is essential for Pol II transcription, but is not a required component of the preinitiation complex in vivo. eLife 6, 155 (2017). This work demonstrates that the depletion of different Mediator subunits affects transcription of a specific subset of genes more strongly than others.

  210. 210.

    Huminiecki, Ł. & Horbańczuk, J. Can we predict gene expression by understanding proximal promoter architecture? Trends Biotechnol. 35, 530–546 (2017).

  211. 211.

    Bonn, S. et al. Cell type-specific chromatin immunoprecipitation from multicellular complex samples using BiTS-ChIP. Nat. Protoc. 7, 978–994 (2012).

  212. 212.

    Lai, W. K. M. & Pugh, B. F. Genome-wide uniformity of human ‘open’ pre-initiation complexes. Genome Res. 27, 15–26 (2017).

  213. 213.

    Andersson, R., Sandelin, A. & Danko, C. G. A unified architecture of transcriptional regulatory elements. Trends Genet. 31, 426–433 (2015).

  214. 214.

    Arnold, C. D. et al. Genome-wide quantitative enhancer activity maps identified by STARR-seq. Science 339, 1074–1077 (2013). This work functionally maps enhancer activity across an entire genome.

  215. 215.

    Muerdter, F. et al. Resolving systematic errors in widely used enhancer activity assays in human cells. Nat. Methods 15, 141–149 (2018).

  216. 216.

    van Arensbergen, J. et al. Genome-wide mapping of autonomous promoter activity in human cells. Nat. Biotechnol. 35, 145–153 (2016). This study reports autonomous promoter activity across the human genome.

  217. 217.

    Nguyen, T. A. et al. High-throughput functional comparison of promoter and enhancer activities. Genome Res. 26, 1023–1033 (2016).

  218. 218.

    Dao, L. T. M. et al. Genome-wide characterization of mammalian promoters with distal enhancer functions. Nat. Genet. 49, 1073–1081 (2017).

  219. 219.

    Catarino, R. R., Neumayr, C. & Stark, A. Promoting transcription over long distances. Nat. Genet. 49, 972–973 (2017).

  220. 220.

    Young, R. S., Kumar, Y., Bickmore, W. A. & Taylor, M. S. Bidirectional transcription initiation marks accessible chromatin and is not specific to enhancers. Genome Biol. 18, 242 (2017).

  221. 221.

    Jin, Y., Eser, U., Struhl, K. & Churchman, L. S. The ground state and evolution of promoter region directionality. Cell 170, 1–21 (2017). This work shows that transcription from newly emerged promoter regions in yeast is bidirectional and that transcription directionality is an evolutionarily selected trait.

  222. 222.

    Neri, F. et al. Intragenic DNA methylation prevents spurious transcription initiation. Nature 543, 72–77 (2017).

  223. 223.

    Kim, J. et al. Blocking promiscuous activation at cryptic promoters directs cell type-specific gene expression. Science 356, 717–721 (2017).

  224. 224.

    Lam, M. T. Y., Li, W., Rosenfeld, M. G. & Glass, C. K. Enhancer RNAs and regulated transcriptional programs. Trends Biochem. Sci. 39, 170–182 (2014).

  225. 225.

    Tsai, A. et al. Nuclear microenvironments modulate transcription from low-affinity enhancers. eLife 6, e1006441 (2017).

  226. 226.

    Hnisz, D., Shrinivas, K., Young, R. A., Chakraborty, A. K. & Sharp, P. A. A. Phase separation model for transcriptional control. Cell 169, 13–23 (2017).

  227. 227.

    Muerdter, F. & Stark, A. Gene regulation: activation through space. Curr. Biol. 26, R895–R898 (2016).

  228. 228.

    Brangwynne, C. P. et al. Germline P granules are liquid droplets that localize by controlled dissolution/condensation. Science 324, 1729–1732 (2009).

  229. 229.

    Kato, M. et al. Cell-free formation of RNA granules: low complexity sequence domains form dynamic fibers within hydrogels. Cell 149, 753–767 (2012).

  230. 230.

    Han, T. W. et al. Cell-free formation of RNA granules: bound RNAs identify features and components of cellular assemblies. Cell 149, 768–779 (2012).

  231. 231.

    Larson, A. G. et al. Liquid droplet formation by HP1α suggests a role for phase separation in heterochromatin. Nature 547, 236–240 (2017).

  232. 232.

    Strom, A. R. et al. Phase separation drives heterochromatin domain formation. Nature 547, 241–245 (2017).

  233. 233.

    Dekker, J. & Mirny, L. The 3D Genome as Moderator of Chromosomal Communication. Cell 164, 1110–1121 (2016).

  234. 234.

    Dixon, J. R., Gorkin, D. U. & Ren, B. Chromatin Domains: The Unit of Chromosome Organization. Mol. Cell 62, 668–680 (2016).

  235. 235.

    Roider, H. G., Lenhard, B., Kanhere, A., Haas, S. A. & Vingron, M. CpG-depleted promoters harbor tissue-specific transcription factor binding signals —implications for motif overrepresentation analyses. Nucleic Acids Res. 37, 6305–6315 (2009).

  236. 236.

    Bernstein, B. E. et al. A bivalent chromatin structure marks key developmental genes in embryonic stem cells. Cell 125, 315–326 (2006).

  237. 237.

    Diao, Y. et al. A tiling-deletion-based genetic screen for cis-regulatory element identification in mammalian cells. Nat. Methods 503, 290–635 (2017).

  238. 238.

    Rajagopal, N. et al. High-throughput mapping of regulatory DNA. Nat. Biotechnol. 34, 167–174 (2016).

  239. 239.

    Lubliner, S. et al. Core promoter sequence in yeast is a major determinant of expression level. Genome Res. 25, 1008–1017 (2015).

  240. 240.

    Patwardhan, R. P. et al. High-resolution analysis of DNA regulatory elements by synthetic saturation mutagenesis. Nat. Biotechnol. 27, 1173–1175 (2009).

  241. 241.

    Bucher, P. Weight matrix descriptions of four eukaryotic RNA polymerase II promoter elements derived from 502 unrelated promoter sequences. J. Mol. Biol. 212, 563–578 (1990).

  242. 242.

    Hahn, S., Buratowski, S., Sharp, P. A. & Guarente, L. Yeast TATA-binding protein TFIID binds to TATA elements with both consensus and nonconsensus DNA sequences. Proc. Natl Acad. Sci. USA 86, 5718–5722 (1989).

  243. 243.

    Arkhipova, I. R. et al. The steps of reverse transcription of Drosophila mobile dispersed genetic elements and U3-R-U5 structure of their LTRs. Cell 44, 555–563 (1986).

  244. 244.

    Li, J. & Gilmour, D. S. Distinct mechanisms of transcriptional pausing orchestrated by GAGA factor and M1BP, a novel transcription factor. EMBO J. 32, 1829–1841 (2013).

  245. 245.

    Hirose, F., Yamaguchi, M., Handa, H., Inomata, Y. & Matsukage, A. Novel 8-base pair sequence (Drosophila DNA replication-related element) and specific binding factor involved in the expression of Drosophila genes for DNA polymerase alpha and proliferating cell nuclear antigen. J. Biol. Chem. 268, 2092–2099 (1993).

  246. 246.

    Parry, T. J. et al. The TCT motif, a key component of an RNA polymerase II transcription system for the translational machinery. Genes Dev. 24, 2013–2018 (2010).

  247. 247.

    Tokusumi, Y., Ma, Y., Song, X., Jacobson, R. H. & Takada, S. The new core promoter element XCPE1 (X Core Promoter Element 1) directs activator-, mediator-, and TATA-binding protein-dependent but TFIID-independent RNA polymerase II transcription from TATA-less promoters. Mol. Cell. Biol. 27, 1844–1858 (2007).

  248. 248.

    Anish, R., Hossain, M. B., Jacobson, R. H. & Takada, S. Characterization of transcription from TATA-less promoters: identification of a new core promoter element XCPE2 and analysis of factor requirements. PLoS ONE 4, e5103 (2009).

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Acknowledgements

The authors thank F. Mürdter, M. A. Zabidi, P. R. Andersen, C. Plaschka and C. Bernecky for helpful comments on the manuscript. V.H. is supported by a long-term postdoctoral fellowship from the Human Frontier Science Program (HFSP, grant number LT000324/2016-L). Research in the Stark group is supported by the European Research Council (ERC) under the European Union’s Horizon 2020 research and innovation programme (grant agreement no. 647320) and by the Austrian Science Fund (FWF, F4303-B09). Basic research at the Institute of Molecular Pathology (IMP), Vienna, Austria, is supported by Boehringer Ingelheim GmbH and the Austrian Research Promotion Agency (FFG).

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Nature Reviews Molecular Cell Biology thanks S. Spicuglia and the other anonymous reviewer(s) for their contribution to the peer review of this work.

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  1. Research Institute of Molecular Pathology (IMP), Vienna Biocenter (VBC), Vienna, Austria

    • Vanja Haberle
    •  & Alexander Stark
  2. Medical University of Vienna, Vienna Biocenter (VBC), Vienna, Austria

    • Alexander Stark

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All authors contributed equally to all aspects of the article.

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The authors declare no competing interests.

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Correspondence to Alexander Stark.

Supplementary information

Glossary

Core promoter

A short sequence flanking the transcription start site that is sufficient to assemble the RNA polymerase II transcription machinery and initiate transcription.

General transcription factors

(GTFs). Proteins that together with RNA polymerase II constitute the transcription machinery at the core promoter.

Enhancers

DNA sequences that contain binding sites for sequence-specific transcription factors and increase the level of transcription from distal core promoters, independently of distance and orientation.

Transcription factors

Proteins that directly bind a specific DNA sequence through their DNA-binding domain and regulate the level of transcription by recruiting RNA polymerase II or transcriptional cofactors through their trans-activation domain.

Transcriptional cofactors

Proteins that do not directly bind DNA but are recruited by DNA-binding transcription factors to regulate transcription of target genes.

Enhancer RNAs

(eRNAs). Short ( <2 kb) unstable non-coding RNAs, usually not spliced or polyadenylated, which are transcribed from enhancers and are rapidly degraded by the exosome.

Nucleosome-depleted region

(NDR). A genomic region depleted of canonical nucleosomes; it is usually associated with active regulatory elements such as promoters and enhancers.

Promoters

Genomic regions encompassing a gene core promoter and an upstream proximal promoter, which together autonomously drive transcription.

Proximal promoter

A transcription-activating sequence immediately upstream of the core promoter (typically up to 250 bp upstream of the transcription start site) that contains binding sites for sequence-specific transcription factors and functions like an enhancer.

Pre-initiation complex

(PIC). A large complex of proteins, including RNA polymerase II and its general transcription factors, that assembles at core promoters and is required for transcription initiation.

CpG islands

(CGIs). GC-rich genomic sequences with a frequency of CpG dinucleotides that is higher than that found in the rest of the genome (which is generally depleted of CpG dinucleotides in mammals).

Piwi-interacting RNA

A type of small non-coding RNA (26–31 nucleotides) that interacts with Argonaute proteins from the Piwi family and mediates transcriptional and post-transcriptional gene silencing of transposable elements.

SAGA complex

SPT–ADA–GCN5-acetyltransferase (SAGA) is a co-activator complex with different chromatin-modifying modules, including, for example, histone acetyltransferase GCN5.

Promoter-proximal pausing

Pausing of RNA polymerase II downstream of the transcription start site; this process controls the transition into productive transcription elongation.

Enhancer responsiveness

The extent to which transcription from a core promoter is induced by a distal enhancer.

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https://doi.org/10.1038/s41580-018-0028-8