Abstract
Ever since microRNAs (miRNAs) were first recognized as an extensive gene family >20 years ago, a broad community of researchers was drawn to investigate the universe of small regulatory RNAs. Although core features of miRNA biogenesis and function were revealed early on, recent years continue to uncover fundamental information on the structural and molecular dynamics of core miRNA machinery, how miRNA substrates and targets are selected from the transcriptome, new avenues for multilevel regulation of miRNA biogenesis and mechanisms for miRNA turnover. Many of these latest insights were enabled by recent technological advances, including massively parallel assays, cryogenic electron microscopy, single-molecule imaging and CRISPR–Cas9 screening. Here, we summarize the current understanding of miRNA biogenesis, function and regulation, and outline challenges to address in the future.
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References
Lee, R. C., Feinbaum, R. L. & Ambros, V. The C. elegans heterochronic gene lin-4 encodes small RNAs with antisense complementarity to lin-14. Cell 75, 843–854 (1993).
Wightman, B., Ha, I. & Ruvkun, G. Posttranscriptional regulation of the heterochronic gene lin-14 by lin-4 mediates temporal pattern formation in C. elegans. Cell 75, 855–862 (1993).
Leviten, M. W., Lai, E. C. & Posakony, J. W. The Drosophila gene Bearded encodes a novel small protein and shares 3′ UTR sequence motifs with multiple Enhancer of split complex genes. Development 124, 4039–4051 (1997).
Lai, E. C. & Posakony, J. W. The Bearded box, a novel 3′ UTR sequence motif, mediates negative post-transcriptional regulation of Bearded and Enhancer of split complex gene expression. Development 124, 4847–4856 (1997).
Lai, E. C., Burks, C. & Posakony, J. W. The K box, a conserved 3′ UTR sequence motif, negatively regulates accumulation of Enhancer of split complex transcripts. Development 125, 4077–4088 (1998).
Lai, E. C. microRNAs are complementary to 3′ UTR sequence motifs that mediate negative post-transcriptional regulation. Nat. Genet. 30, 363–364 (2002).
Lau, N., Lim, L., Weinstein, E. & Bartel, D. P. An abundant class of tiny RNAs with probable regulatory roles in Caenorhabditis elegans. Science 294, 858–862 (2001).
Lee, R. C. & Ambros, V. An extensive class of small RNAs in Caenorhabditis elegans. Science 294, 862–864 (2001).
Lagos-Quintana, M., Rauhut, R., Lendeckel, W. & Tuschl, T. Identification of novel genes coding for small expressed RNAs. Science 294, 853–858 (2001).
Bartel, D. P. Metazoan microRNAs. Cell 173, 20–51 (2018).
Czech, B. & Hannon, G. J. Small RNA sorting: matchmaking for Argonautes. Nat. Rev. Genet. 12, 19–31 (2010).
Yang, J. S. & Lai, E. C. Alternative miRNA biogenesis pathways and the interpretation of core miRNA pathway mutants. Mol. Cell 43, 892–903 (2011).
Maurin, T., Cazalla, D., Yang, J. S., Bortolamiol-Becet, D. & Lai, E. C. RNase III-independent microRNA biogenesis in mammalian cells. RNA 18, 2166–2173 (2012).
Shang, R. et al. Ribozyme-enhanced single-stranded Ago2-processed interfering RNA triggers efficient gene silencing with fewer off-target effects. Nat. Commun. 6, 8430 (2015).
Lai, E. C., Tam, B. & Rubin, G. M. Pervasive regulation of Drosophila Notch target genes by GY-box-, Brd-box-, and K-box-class microRNAs. Genes Dev. 19, 1067–1080 (2005).
Lewis, B. P., Burge, C. B. & Bartel, D. P. Conserved seed pairing, often flanked by adenosines, indicates that thousands of human genes are microRNA targets. Cell 120, 15–20 (2005).
Brennecke, J., Stark, A., Russell, R. B. & Cohen, S. M. Principles of microRNA–target recognition. PLoS Biol. 3, e85 (2005).
Wang, Y., Sheng, G., Juranek, S., Tuschl, T. & Patel, D. J. Structure of the guide-strand-containing Argonaute silencing complex. Nature 456, 209–213 (2008).
Schirle, N. T. & MacRae, I. J. The crystal structure of human Argonaute2. Science 336, 1037–1040 (2012).
Elkayam, E. et al. The structure of human Argonaute-2 in complex with miR-20a. Cell 150, 100–110 (2012).
Liu, J. et al. Argonaute2 is the catalytic engine of mammalian RNAi. Science 305, 1437–1441 (2004).
Meister, G. et al. Human Argonaute2 mediates RNA cleavage targeted by miRNAs and siRNAs. Mol. Cell 15, 185–197 (2004).
Park, M. S. et al. Human Argonaute3 has slicer activity. Nucleic acids Res. 45, 11867–11877 (2017).
Park, M. S., Sim, G., Kehling, A. C. & Nakanishi, K. Human Argonaute2 and Argonaute3 are catalytically activated by different lengths of guide RNA. Proc. Natl Acad. Sci. USA 117, 28576–28578 (2020).
Nguyen, H. M., Nguyen, T. D., Nguyen, T. L. & Nguyen, T. A. Orientation of human microprocessor on primary microRNAs. Biochemistry 58, 189–198 (2019).
Han, J. et al. Molecular basis for the recognition of primary microRNAs by the Drosha–DGCR8 complex. Cell 125, 887–901 (2006).
Zeng, Y. & Cullen, B. R. Efficient processing of primary microRNA hairpins by Drosha requires flanking non-structured RNA sequences. J. Biol. Chem. 280, 27595–27603 (2005).
Zeng, Y., Yi, R. & Cullen, B. R. Recognition and cleavage of primary microRNA precursors by the nuclear processing enzyme Drosha. EMBO J. 24, 138–148 (2005).
Ma, H., Wu, Y., Choi, J. G. & Wu, H. Lower and upper stem-single-stranded RNA junctions together determine the Drosha cleavage site. Proc. Natl Acad. Sci. USA 110, 20687–20692 (2013).
Auyeung, V. C., Ulitsky, I., McGeary, S. E. & Bartel, D. P. Beyond secondary structure: primary-sequence determinants license pri-miRNA hairpins for processing. Cell 152, 844–858 (2013). This study is the first to utilize massively parallel substrate assays to reveal motifs involved in pri-miRNA processing.
Fang, W. & Bartel, D. P. The menu of features that define primary microRNAs and enable de novo design of microRNA genes. Mol. Cell 60, 131–145 (2015).
Partin, A. C. et al. Heme enables proper positioning of Drosha and DGCR8 on primary microRNAs. Nat. Commun. 8, 1737 (2017).
Nguyen, T. A., Park, J., Dang, T. L., Choi, Y. G. & Kim, V. N. Microprocessor depends on hemin to recognize the apical loop of primary microRNA. Nucleic acids Res. 46, 5726–5736 (2018).
Kim, K., Nguyen, T. D., Li, S. & Nguyen, T. A. SRSF3 recruits DROSHA to the basal junction of primary microRNAs. RNA 24, 892–898 (2018).
Fernandez, N. et al. Genetic variation and RNA structure regulate microRNA biogenesis. Nat. Commun. 8, 15114 (2017).
Li, S., Nguyen, T. D., Nguyen, T. L. & Nguyen, T. A. Mismatched and wobble base pairs govern primary microRNA processing by human microprocessor. Nat. Commun. 11, 1926 (2020).
Li, S., Le, T. N., Nguyen, T. D., Trinh, T. A. & Nguyen, T. A. Bulges control pri-miRNA processing in a position and strand-dependent manner. RNA Biol. 18, 1716–1726 (2021).
Lee, Y. Y., Kim, H. & Kim, V. N. Sequence determinant of small RNA production by DICER. Nature 615, 323–330 (2023).
Rice, G. M., Shivashankar, V., Ma, E. J., Baryza, J. L. & Nutiu, R. Functional atlas of primary miRNA maturation by the microprocessor. Mol. Cell 80, 892–902.e4 (2020).
Kim, K. et al. A quantitative map of human primary microRNA processing sites. Mol. Cell 81, 3422–3439.e11 (2021).
Kang, W. et al. MapToCleave: high-throughput profiling of microRNA biogenesis in living cells. Cell Rep. 37, 110015 (2021).
Chiang, H. R. et al. Mammalian microRNAs: experimental evaluation of novel and previously annotated genes. Genes Dev. 24, 992–1009 (2010).
Consortium, R. RNAcentral 2021: secondary structure integration, improved sequence search and new member databases. Nucleic acids Res. 49, D212–D220 (2021).
Fromm, B., Zhong, X., Tarbier, M., Friedlander, M. R. & Hackenberg, M. The limits of human microRNA annotation have been met. RNA https://doi.org/10.1261/rna.079098.122 (2022).
Nguyen, T. L., Nguyen, T. D., Bao, S., Li, S. & Nguyen, T. A. The internal loops in the lower stem of primary microRNA transcripts facilitate single cleavage of human microprocessor. Nucleic acids Res. 48, 2579–2593 (2020).
Luo, Q. J. et al. RNA structure probing reveals the structural basis of Dicer binding and cleavage. Nat. Commun. 12, 3397 (2021).
Zhang, H., Kolb, F. A., Jaskiewicz, L., Westhof, E. & Filipowicz, W. Single processing center models for human Dicer and bacterial RNase III. Cell 118, 57–68 (2004).
Macrae, I. J. et al. Structural basis for double-stranded RNA processing by Dicer. Science 311, 195–198 (2006).
MacRae, I. J., Zhou, K. & Doudna, J. A. Structural determinants of RNA recognition and cleavage by Dicer. Nat. Struct. Mol. Biol. 14, 934–940 (2007).
Vermeulen, A. et al. The contributions of dsRNA structure to Dicer specificity and efficiency. RNA 11, 674–682 (2005).
Zhang, H., Kolb, F. A., Brondani, V., Billy, E. & Filipowicz, W. Human Dicer preferentially cleaves dsRNAs at their termini without a requirement for ATP. EMBO J. 21, 5875–5885 (2002).
Park, J. E. et al. Dicer recognizes the 5′ end of RNA for efficient and accurate processing. Nature 475, 201–205 (2011).
Gu, S. et al. The loop position of shRNAs and pre-miRNAs is critical for the accuracy of dicer processing in vivo. Cell 151, 900–911 (2012).
Nguyen, T. D., Trinh, T. A., Bao, S. & Nguyen, T. A. Secondary structure RNA elements control the cleavage activity of DICER. Nat. Commun. 13, 2138 (2022).
Frank, F., Sonenberg, N. & Nagar, B. Structural basis for 5′-nucleotide base-specific recognition of guide RNA by human AGO2. Nature 465, 818–822 (2010).
Boland, A., Tritschler, F., Heimstadt, S., Izaurralde, E. & Weichenrieder, O. Crystal structure and ligand binding of the MID domain of a eukaryotic Argonaute protein. EMBO Rep. 11, 522–527 (2010).
Schirle, N. T., Sheu-Gruttadauria, J., Chandradoss, S. D., Joo, C. & MacRae, I. J. Water-mediated recognition of t1-adenosine anchors Argonaute2 to microRNA targets. eLife https://doi.org/10.7554/eLife.07646 (2015).
Shin, C. et al. Expanding the microRNA targeting code: functional sites with centered pairing. Mol. Cell 38, 789–802 (2010).
Chi, S. W., Hannon, G. J. & Darnell, R. B. An alternative mode of microRNA target recognition. Nat. Struct. Mol. Biol. 19, 321–327 (2012).
Lal, A. et al. miR-24 Inhibits cell proliferation by targeting E2F2, MYC, and other cell-cycle genes via binding to “seedless” 3′UTR microRNA recognition elements. Mol. Cell 35, 610–625 (2009).
Loeb, G. B. et al. Transcriptome-wide miR-155 binding map reveals widespread noncanonical microRNA targeting. Mol. Cell 48, 760–770 (2012).
Hafner, M. et al. Transcriptome-wide identification of RNA-binding protein and microRNA target sites by PAR-CLIP. Cell 141, 129–141 (2010).
Becker, W. R. et al. High-throughput analysis reveals rules for target RNA binding and cleavage by AGO2. Mol. Cell 75, 741–755.e11 (2019).
McGeary, S. E. et al. The biochemical basis of microRNA targeting efficacy. Science https://doi.org/10.1126/science.aav1741 (2019). This paper reveals broad features of miRNA–target interactions using RBNS.
McGeary, S. E., Bisaria, N., Pham, T. M., Wang, P. Y. & Bartel, D. P. microRNA 3′-compensatory pairing occurs through two binding modes, with affinity shaped by nucleotide identity and position. eLife https://doi.org/10.7554/eLife.69803 (2022).
Grimson, A. et al. microRNA targeting specificity in mammals: determinants beyond seed pairing. Mol. Cell 27, 91–105 (2007).
Gu, S., Jin, L., Zhang, F., Sarnow, P. & Kay, M. A. Biological basis for restriction of microRNA targets to the 3′ untranslated region in mammalian mRNAs. Nat. Struct. Mol. Biol. 16, 144–150 (2009).
Schnall-Levin, M. et al. Unusually effective microRNA targeting within repeat-rich coding regions of mammalian mRNAs. Genome Res. 21, 1395–1403 (2011).
Zhang, K. et al. A novel class of microRNA-recognition elements that function only within open reading frames. Nat. Struct. Mol. Biol. 25, 1019–1027 (2018).
Yu, S. & Kim, V. N. A tale of non-canonical tails: gene regulation by post-transcriptional RNA tailing. Nat. Rev. Mol. Cell Biol. https://doi.org/10.1038/s41580-020-0246-8 (2020).
Yang, A. et al. 3′ uridylation confers miRNAs with non-canonical target repertoires. Mol. Cell 75, 511–522.e4 (2019).
Sheu-Gruttadauria, J., Xiao, Y., Gebert, L. F. & MacRae, I. J. Beyond the seed: structural basis for supplementary microRNA targeting by human Argonaute2. EMBO J. 38, e101153 (2019).
Sim, G. et al. Manganese-dependent microRNA trimming by 3′ → 5′ exonucleases generates 14-nucleotide or shorter tiny RNAs. Proc. Natl Acad. Sci. USA 119, e2214335119 (2022).
Kim, H. H. et al. HuR recruits let-7/RISC to repress c-Myc expression. Genes Dev. 23, 1743–1748 (2009).
Sternburg, E. L., Estep, J. A., Nguyen, D. K., Li, Y. & Karginov, F. V. Antagonistic and cooperative AGO2–PUM interactions in regulating mRNAs. Sci. Rep. 8, 15316 (2018).
Kedde, M. et al. RNA-binding protein Dnd1 inhibits microRNA access to target mRNA. Cell 131, 1273–1286 (2007).
Kim, S. et al. The regulatory impact of RNA-binding proteins on microRNA targeting. Nat. Commun. 12, 5057 (2021).
Wang, Y. et al. Structure of an argonaute silencing complex with a seed-containing guide DNA and target RNA duplex. Nature 456, 921–926 (2008).
Wang, Y. et al. Nucleation, propagation and cleavage of target RNAs in Ago silencing complexes. Nature 461, 754–761 (2009).
Song, J. J., Smith, S. K., Hannon, G. J. & Joshua-Tor, L. Crystal structure of Argonaute and its implications for RISC slicer activity. Science 305, 1434–1437 (2004).
Liu, Z. et al. Cryo-EM structure of human dicer and its complexes with a pre-miRNA substrate. Cell 173, 1191–1203.e12 (2018).
Nguyen, T. A. et al. Functional anatomy of the human microprocessor. Cell 161, 1374–1387 (2015).
Kwon, S. C. et al. Structure of human DROSHA. Cell 164, 81–90 (2016).
Barr, I. et al. DiGeorge critical region 8 (DGCR8) is a double-cysteine-ligated heme protein. J. Biol. Chem. 286, 16716–16725 (2011).
Quick-Cleveland, J. et al. The DGCR8 RNA-binding heme domain recognizes primary microRNAs by clamping the hairpin. Cell Rep. 7, 1994–2005 (2014).
Jin, W., Wang, J., Liu, C. P., Wang, H. W. & Xu, R. M. Structural basis for pri-miRNA recognition by Drosha. Mol. Cell 78, 423–433.e5 (2020).
Partin, A. C. et al. Cryo-EM structures of human Drosha and DGCR8 in complex with primary microRNA. Mol. Cell 78, 411–422 e414 (2020). Together with Jin et al. (2020), this work is the first cryo-EM study of Microprocessor structures.
Shi, Z., Nicholson, R. H., Jaggi, R. & Nicholson, A. W. Characterization of Aquifex aeolicus ribonuclease III and the reactivity epitopes of its pre-ribosomal RNA substrates. Nucleic acids Res. 39, 2756–2768 (2011).
Carmell, M. A. & Hannon, G. J. RNase III enzymes and the initiation of gene silencing. Nat. Struct. Mol. Biol. 11, 214–218 (2004).
Lee, Y. Y., Lee, H., Kim, H., Kim, V. N. & Roh, S. H. Structure of the human DICER–pre-miRNA complex in a dicing state. Nature 615, 331–338 (2023).
Zapletal, D. et al. Structural and functional basis of mammalian microRNA biogenesis by Dicer. Mol. Cell 82, 4064–4079.e13 (2022). Together with Lee et al. (Nature, 2023), this work reports new cryo-EM structures for active mammalian Dicer complex.
Jouravleva, K. et al. Structural basis of microRNA biogenesis by Dicer-1 and its partner protein Loqs-PB. Mol. Cell 82, 4049–4063.e6 (2022).
Wei, X. et al. Structural basis of microRNA processing by Dicer-like 1. Nat. Plants 7, 1389–1396 (2021).
Fareh, M. et al. TRBP ensures efficient Dicer processing of precursor microRNA in RNA-crowded environments. Nat. Commun. 7, 13694 (2016).
Yamaguchi, S. et al. Structure of the Dicer-2–R2D2 heterodimer bound to a small RNA duplex. Nature 607, 393–398 (2022).
Su, S. et al. Structural insights into dsRNA processing by Drosophila Dicer-2–Loqs-PD. Nature 607, 399–406 (2022).
Wang, Q. et al. Mechanism of siRNA production by a plant Dicer–RNA complex in dicing-competent conformation. Science 374, 1152–1157 (2021).
Herbert, K. M. et al. A heterotrimer model of the complete Microprocessor complex revealed by single-molecule subunit counting. RNA 22, 175–183 (2016).
Naganuma, M., Tadakuma, H. & Tomari, Y. Single-molecule analysis of processive double-stranded RNA cleavage by Drosophila Dicer-2. Nat. Commun. 12, 4268 (2021).
Iwasaki, S. et al. Defining fundamental steps in the assembly of the Drosophila RNAi enzyme complex. Nature 521, 533–536 (2015).
Tsuboyama, K., Tadakuma, H. & Tomari, Y. Conformational activation of argonaute by distinct yet coordinated actions of the Hsp70 and Hsp90 chaperone systems. Mol. Cell 70, 722–729.e4 (2018).
Willkomm, S. et al. Single-molecule FRET uncovers hidden conformations and dynamics of human Argonaute 2. Nat. Commun. 13, 3825 (2022). This study uses single-molecule fluorescence to dissect internal motions within human Ago2 during transitions from guide RNA binding to target capture.
Salomon, W. E., Jolly, S. M., Moore, M. J., Zamore, P. D. & Serebrov, V. Single-molecule imaging reveals that argonaute reshapes the binding properties of its nucleic acid guides. Cell 162, 84–95 (2015).
Yao, C., Sasaki, H. M., Ueda, T., Tomari, Y. & Tadakuma, H. Single-molecule analysis of the target cleavage reaction by the Drosophila RNAi enzyme complex. Mol. Cell 59, 125–132 (2015).
Chandradoss, S. D., Schirle, N. T., Szczepaniak, M., MacRae, I. J. & Joo, C. A dynamic search process underlies microRNA targeting. Cell 162, 96–107 (2015). This study uses the distance sensing capability of smFRET to visualize strategies of target interrogation by RISC.
Klum, S. M., Chandradoss, S. D., Schirle, N. T., Joo, C. & MacRae, I. J. Helix-7 in argonaute2 shapes the microRNA seed region for rapid target recognition. EMBO J. 37, 75–88 (2018).
Cui, T. J. et al. Argonaute bypasses cellular obstacles without hindrance during target search. Nat. Commun. 10, 4390 (2019).
Ruijtenberg, S. et al. mRNA structural dynamics shape Argonaute–target interactions. Nat. Struct. Mol. Biol. 27, 790–801 (2020).
Cialek, C. A. et al. Imaging translational control by Argonaute with single-molecule resolution in live cells. Nat. Commun. 13, 3345 (2022).
Kobayashi, H. & Singer, R. H. Single-molecule imaging of microRNA-mediated gene silencing in cells. Nat. Commun. 13, 1435 (2022).
Sheu-Gruttadauria, J. & MacRae, I. J. Structural foundations of RNA silencing by Argonaute. J. Mol. Biol. 429, 2619–2639 (2017).
Nakanishi, K. Anatomy of four human Argonaute proteins. Nucleic acids Res. 50, 6618–6638 (2022).
Sheu-Gruttadauria, J. et al. Structural basis for target-directed microRNA degradation. Mol. Cell https://doi.org/10.1016/j.molcel.2019.06.019 (2019).
Newman, M. A., Thomson, J. M. & Hammond, S. M. Lin-28 interaction with the Let-7 precursor loop mediates regulated microRNA processing. RNA 14, 1539–1549 (2008).
Heo, I. et al. Lin28 mediates the terminal uridylation of let-7 precursor microRNA. Mol. Cell 32, 276–284 (2008).
Viswanathan, S. R., Daley, G. Q. & Gregory, R. I. Selective blockade of microRNA processing by Lin28. Science 320, 97–100 (2008).
Rybak, A. et al. A feedback loop comprising lin-28 and let-7 controls pre-let-7 maturation during neural stem-cell commitment. Nat. Cell Biol. 10, 987–993 (2008).
Heo, I. et al. TUT4 in concert with Lin28 suppresses microRNA biogenesis through pre-microRNA uridylation. Cell 138, 696–708 (2009).
Thornton, J. E., Chang, H. M., Piskounova, E. & Gregory, R. I. Lin28-mediated control of let-7 microRNA expression by alternative TUTases Zcchc11 (TUT4) and Zcchc6 (TUT7). RNA 18, 1875–1885 (2012).
Chang, H. M., Triboulet, R., Thornton, J. E. & Gregory, R. I. A role for the Perlman syndrome exonuclease Dis3l2 in the Lin28–let-7 pathway. Nature https://doi.org/10.1038/nature12119 (2013).
Faehnle, C. R., Walleshauser, J. & Joshua-Tor, L. Mechanism of Dis3l2 substrate recognition in the Lin28–let-7 pathway. Nature 514, 252–256 (2014).
Ustianenko, D. et al. Mammalian DIS3L2 exoribonuclease targets the uridylated precursors of let-7 miRNAs. RNA 19, 1632–1638 (2013).
Heo, I. et al. Mono-uridylation of pre-microRNA as a key step in the biogenesis of group II let-7 microRNAs. Cell 151, 521–532 (2012).
Kim, H. et al. A mechanism for microRNA arm switching regulated by uridylation. Mol. Cell 78, 1224–1236.e5 (2020).
Michlewski, G. & Caceres, J. F. Post-transcriptional control of miRNA biogenesis. RNA 25, 1–16 (2019).
Treiber, T., Treiber, N. & Meister, G. Regulation of microRNA biogenesis and its crosstalk with other cellular pathways. Nat. Rev. Mol. Cell Biol. 20, 5–20 (2019).
Michlewski, G., Guil, S., Semple, C. A. & Caceres, J. F. Posttranscriptional regulation of miRNAs harboring conserved terminal loops. Mol. Cell 32, 383–393 (2008).
Nussbacher, J. K. & Yeo, G. W. Systematic discovery of RNA binding proteins that regulate microRNA levels. Mol. Cell 69, 1005–1016.e7 (2018).
Treiber, T. et al. A compendium of RNA-binding proteins that regulate microRNA biogenesis. Mol. Cell 66, 270–284.e13 (2017).
Tokumaru, S., Suzuki, M., Yamada, H., Nagino, M. & Takahashi, T. let-7 regulates Dicer expression and constitutes a negative feedback loop. Carcinogenesis 29, 2073–2077 (2008).
Martello, G. et al. A microRNA targeting dicer for metastasis control. Cell 141, 1195–1207 (2010).
Wang, D. et al. Uncovering the cellular capacity for intensive and specific feedback self-control of the argonautes and microRNA targeting activity. Nucleic acids Res. 48, 4681–4697 (2020).
Smibert, P., Yang, J. S., Azzam, G., Liu, J. L. & Lai, E. C. Homeostatic control of Argonaute stability by microRNA availability. Nat. Struct. Mol. Biol. 20, 789–795 (2013).
Kobayashi, H., Shoji, K., Kiyokawa, K., Negishi, L. & Tomari, Y. Iruka eliminates dysfunctional argonaute by selective ubiquitination of its empty state. Mol. Cell 73, 119–129.e5 (2019).
Martinez, N. J. & Gregory, R. I. Argonaute2 expression is post-transcriptionally coupled to microRNA abundance. RNA 19, 605–612 (2013).
Derrien, B. et al. Degradation of the antiviral component ARGONAUTE1 by the autophagy pathway. Proc. Natl Acad. Sci. USA 109, 15942–15946 (2012).
Han, J. et al. Posttranscriptional crossregulation between Drosha and DGCR8. Cell 136, 75–84 (2009). This paper identifies the first cross-regulations between miRNA factors, in this case between Microprocessor components Drosha and DGCR8 (see also references 138 and 139).
Kadener, S. et al. Genome-wide identification of targets of the Drosha-Pasha/DGCR8 complex. RNA 15, 537–545 (2009).
Smibert, P. et al. A Drosophila genetic screen yields allelic series of core microRNA biogenesis factors and reveals post-developmental roles for microRNAs. RNA 17, 1997–2010 (2011).
Cui, Y. et al. Global miRNA dosage control of embryonic germ layer specification. Nature 593, 602–606 (2021).
Baskerville, S. & Bartel, D. P. Microarray profiling of microRNAs reveals frequent coexpression with neighboring miRNAs and host genes. RNA 11, 241–247 (2005).
Du, P., Wang, L., Sliz, P. & Gregory, R. I. A biogenesis step upstream of microprocessor controls miR-17 approximately 92 expression. Cell 162, 885–899 (2015).
Donayo, A. O. et al. Oncogenic biogenesis of pri-miR-17-92 reveals hierarchy and competition among polycistronic microRNAs. Mol. Cell 75, 340–356 (2019).
Truscott, M., Islam, A. B. & Frolov, M. V. Novel regulation and functional interaction of polycistronic miRNAs. RNA 22, 129–138 (2016).
Lataniotis, L. et al. CRISPR/Cas9 editing reveals novel mechanisms of clustered microRNA regulation and function. Sci. Rep. 7, 8585 (2017).
Haar, J. et al. The expression of a viral microRNA is regulated by clustering to allow optimal B cell transformation. Nucleic acids Res. 44, 1326–1341 (2016).
Vilimova, M. et al. Cis regulation within a cluster of viral microRNAs. Nucleic acids Res. 49, 10018–10033 (2021).
Yang, J. S. & Lai, E. C. Dicer-independent, Ago2-mediated microRNA biogenesis in vertebrates. Cell Cycle 9, 4455–4460 (2010).
Jee, D. et al. Dual strategies for Argonaute2-mediated biogenesis of erythroid miRNAs underlie conserved requirements for slicing in mammals. Mol. Cell 69, 265–278.e6 (2018).
Kretov, D. A. et al. Ago2-dependent processing allows miR-451 to evade the global microRNA turnover elicited during erythropoiesis. Mol. Cell 78, 317–328.e6 (2020).
Shang, R. et al. Genomic clustering facilitates nuclear processing of suboptimal pri-miRNA loci. Mol. Cell 78, 303–316.e4 (2020).
Fang, W. & Bartel, D. P. microRNA clustering assists processing of suboptimal microRNA hairpins through the action of the ERH protein. Mol. Cell 78, 289–302.e6 (2020).
Hutter, K. et al. SAFB2 enables the processing of suboptimal stem–loop structures in clustered primary miRNA transcripts. Mol. Cell 78, 876–889 (2020). Together with Shang et al. (2020) and Fang and Bartel (2020), this work reports mechanisms for miRNA cluster assistance for suboptimal miRNA biogenesis.
Kwon, S. C. et al. ERH facilitates microRNA maturation through the interaction with the N-terminus of DGCR8. Nucleic acids Res. 48, 11097–11112 (2020).
Kim, Y. K. & Kim, V. N. Processing of intronic microRNAs. EMBO J. 26, 775–783 (2007).
Ballarino, M. et al. Coupled RNA processing and transcription of intergenic primary microRNAs. Mol. Cell. Biol. 29, 5632–5638 (2009).
Morlando, M. et al. Primary microRNA transcripts are processed co-transcriptionally. Nat. Struct. Mol. Biol. 15, 902–909 (2008).
Liu, H. et al. HP1BP3, a chromatin retention factor for co-transcriptional microRNA processing. Mol. Cell 63, 420–432 (2016).
Church, V. A. et al. Microprocessor recruitment to elongating RNA polymerase II is required for differential expression of microRNAs. Cell Rep. 20, 3123–3134 (2017).
Pawlicki, J. M. & Steitz, J. A. Primary microRNA transcript retention at sites of transcription leads to enhanced microRNA production. J. Cell Biol. 182, 61–76 (2008).
Bernstein, E., Caudy, A., Hammond, S. & Hannon, G. Role for a bidentate ribonuclease in the initiation step of RNA interference. Nature 409, 363–366 (2001).
Hutvagner, G. et al. A cellular function for the RNA-interference enzyme Dicer in the maturation of the let-7 small temporal RNA. Science 293, 834–838 (2001).
Flemr, M. et al. A retrotransposon-driven dicer isoform directs endogenous small interfering RNA production in mouse oocytes. Cell 155, 807–816 (2013). This paper reports that an endogenous N-terminally truncated Dicer isoform processes siRNAs in mouse oocytes, an unusual setting for mammalian siRNA biology.
Watanabe, T. et al. Endogenous siRNAs from naturally formed dsRNAs regulate transcripts in mouse oocytes. Nature 453, 539–543 (2008).
Tam, O. H. et al. Pseudogene-derived small interfering RNAs regulate gene expression in mouse oocytes. Nature 453, 534–538 (2008).
Suh, N. et al. microRNA function is globally suppressed in mouse oocytes and early embryos. Curr. Biol. 20, 271–277 (2010).
Murchison, E. P. et al. Critical roles for Dicer in the female germline. Genes Dev. 21, 682–693 (2007).
Kataruka, S. et al. microRNA dilution during oocyte growth disables the microRNA pathway in mammalian oocytes. Nucleic acids Res. 48, 8050–8062 (2020).
Schuster, S., Miesen, P. & van Rij, R. P. Antiviral RNAi in insects and mammals: parallels and differences. Viruses https://doi.org/10.3390/v11050448 (2019).
Kennedy, E. M. et al. Production of functional small interfering RNAs by an amino-terminal deletion mutant of human Dicer. Proc. Natl Acad. Sci. USA 112, E6945–E6954 (2015).
Poirier, E. Z. et al. An isoform of Dicer protects mammalian stem cells against multiple RNA viruses. Science 373, 231–236 (2021).
Li, Y., Lu, J., Han, Y., Fan, X. & Ding, S. W. RNA interference functions as an antiviral immunity mechanism in mammals. Science 342, 231–234 (2013).
Maillard, P. V. et al. Antiviral RNA interference in mammalian cells. Science 342, 235–238 (2013).
Gantier, M. P. et al. Analysis of microRNA turnover in mammalian cells following Dicer1 ablation. Nucleic acids Res. 39, 5692–5703 (2011).
van Rooij, E. et al. Control of stress-dependent cardiac growth and gene expression by a microRNA. Science 316, 575–579 (2007).
Kingston, E. R. & Bartel, D. P. Global analyses of the dynamics of mammalian microRNA metabolism. Genome Res. 29, 1777–1790 (2019).
Gibbings, D. et al. Selective autophagy degrades DICER and AGO2 and regulates miRNA activity. Nat. Cell Biol. 14, 1314–1321 (2012).
Reichholf, B. et al. Time-resolved small RNA sequencing unravels the molecular principles of microRNA homeostasis. Mol. Cell 75, 756–768.e7 (2019).
Krol, J. et al. Characterizing light-regulated retinal microRNAs reveals rapid turnover as a common property of neuronal microRNAs. Cell 141, 618–631 (2010).
Ramachandran, V. & Chen, X. Degradation of microRNAs by a family of exoribonucleases in Arabidopsis. Science 321, 1490–1492 (2008).
Shukla, S., Bjerke, G. A., Muhlrad, D., Yi, R. & Parker, R. The RNase PARN controls the levels of specific miRNAs that contribute to p53 regulation. Mol. Cell 73, 1204–1216.e4 (2019).
Yu, Y. et al. ARGONAUTE10 promotes the degradation of miR165/6 through the SDN1 and SDN2 exonucleases in Arabidopsis. PLoS Biol. 15, e2001272 (2017).
Ibrahim, F. et al. Uridylation of mature miRNAs and siRNAs by the MUT68 nucleotidyltransferase promotes their degradation in Chlamydomonas. Proc. Natl Acad. Sci. USA 107, 3906–3911 (2010).
Chatterjee, S. & Grosshans, H. Active turnover modulates mature microRNA activity in Caenorhabditis elegans. Nature 461, 546–549 (2009).
Cazalla, D., Yario, T. & Steitz, J. A. Down-regulation of a host microRNA by a Herpesvirus saimiri noncoding RNA. Science 328, 1563–1566 (2010).
Marcinowski, L. et al. Degradation of cellular mir-27 by a novel, highly abundant viral transcript is important for efficient virus replication in vivo. PLoS Pathog. 8, e1002510 (2012).
Ghini, F. et al. Endogenous transcripts control miRNA levels and activity in mammalian cells by target-directed miRNA degradation. Nat. Commun. 9, 3119 (2018).
de la Mata, M. et al. Potent degradation of neuronal miRNAs induced by highly complementary targets. EMBO Rep. https://doi.org/10.15252/embr.201540078 (2015).
Ameres, S. L. et al. Target RNA-directed trimming and tailing of small silencing RNAs. Science 328, 1534–1539 (2010).
Ameres, S. L. & Zamore, P. D. Diversifying microRNA sequence and function. Nat. Rev. Mol. Cell Biol. 14, 475–488 (2013).
Yang, A. et al. AGO-bound mature miRNAs are oligouridylated by TUTs and subsequently degraded by DIS3L2. Nat. Commun. 11, 2765 (2020).
Yang, A. et al. TENT2, TUT4, and TUT7 selectively regulate miRNA sequence and abundance. Nat. Commun. 13, 5260 (2022).
Shi, C. Y. et al. The ZSWIM8 ubiquitin ligase mediates target-directed microRNA degradation. Science https://doi.org/10.1126/science.abc9359 (2020).
Han, J. et al. A ubiquitin ligase mediates target-directed microRNA decay independently of tailing and trimming. Science https://doi.org/10.1126/science.abc9546 (2020). Together with Shi et al. (2020), this work reveals the regulatory mechanism of TDMD, via ZSWIM8-mediated Argonaute protein degradation.
Piwecka, M. et al. Loss of a mammalian circular RNA locus causes miRNA deregulation and affects brain function. Science https://doi.org/10.1126/science.aam8526 (2017).
Ulitsky, I., Shkumatava, A., Jan, C. H., Sive, H. & Bartel, D. P. Conserved function of lincRNAs in vertebrate embryonic development despite rapid sequence evolution. Cell 147, 1537–1550 (2011).
Kleaveland, B., Shi, C. Y., Stefano, J. & Bartel, D. P. A network of noncoding regulatory RNAs acts in the mammalian brain. Cell 174, 350–362.e17 (2018).
Donnelly, B. F. et al. The developmentally timed decay of an essential microRNA family is seed-sequence dependent. Cell Rep. 40, 111154 (2022).
Li, L. et al. Widespread microRNA degradation elements in target mRNAs can assist the encoded proteins. Genes Dev. 35, 1595–1609 (2021).
Bitetti, A. et al. microRNA degradation by a conserved target RNA regulates animal behavior. Nat. Struct. Mol. Biol. 25, 244–251 (2018).
Kingston, E. R., Blodgett, L. W. & Bartel, D. P. Endogenous transcripts direct microRNA degradation in Drosophila, and this targeted degradation is required for proper embryonic development. Mol. Cell 82, 3872–3884.e9 (2022).
Sheng, P. et al. Screening of Drosophila microRNA-degradation sequences reveals Argonaute1 mRNA’s role in regulating miR-999. Nat. Commun. 14, 2108 (2023).
Katoh, T. et al. Selective stabilization of mammalian microRNAs by 3′ adenylation mediated by the cytoplasmic poly(A) polymerase GLD-2. Genes Dev. 23, 433–438 (2009).
D’Ambrogio, A., Gu, W., Udagawa, T., Mello, C. C. & Richter, J. D. Specific miRNA stabilization by Gld2-catalyzed monoadenylation. Cell Rep. https://doi.org/10.1016/j.celrep.2012.10.023 (2012).
Jeong, H. C. et al. USB1 is a miRNA deadenylase that regulates hematopoietic development. Science 379, 901–907 (2023).
Mansur, F. et al. Gld2-catalyzed 3′ monoadenylation of miRNAs in the hippocampus has no detectable effect on their stability or on animal behavior. RNA 22, 1492–1499 (2016).
Vieux, K. F. et al. Screening by deep sequencing reveals mediators of microRNA tailing in C. elegans. Nucleic Acids Res. 49, 11167–11180 (2021).
Lee, S. et al. Promiscuous splicing-derived hairpins are dominant substrates of tailing-mediated defense of miRNA biogenesis in mammals. Cell Rep. 42, 112111 (2023).
Nishida, K. M. et al. Roles of R2D2, a cytoplasmic D2 body component, in the endogenous siRNA pathway in Drosophila. Mol. Cell 49, 680–691 (2013).
Drake, M. et al. A requirement for ERK dependent Dicer phosphorylation in coordinating oocyte-to-embryo transition in Caenorhabditis elegans. Dev. Cell 31, 614–628 (2014).
Shang, R. et al. Regulated dicing of pre-mir-144 via reshaping of its terminal loop. Nucleic acids Res. 50, 7637–7654 (2022).
Gutierrez-Perez, P. et al. miR-1 sustains muscle physiology by controlling V-ATPase complex assembly. Sci. Adv. 7, eabh1434 (2021).
Yang, B., Schwartz, M. & McJunkin, K. In vivo CRISPR screening for phenotypic targets of the mir-35-42 family in C. elegans. Genes Dev. 34, 1227–1238 (2020).
Ecsedi, M., Rausch, M. & Grosshans, H. The let-7 microRNA directs vulval development through a single target. Dev. Cell 32, 335–344 (2015).
Garaulet, D. L., Zhang, B., Wei, L., Li, E. & Lai, E. C. miRNAs and neural alternative polyadenylation specify the virgin behavioral state. Dev. Cell 54, 410–423 (2020).
Okamura, K., Hagen, J. W., Duan, H., Tyler, D. M. & Lai, E. C. The mirtron pathway generates microRNA-class regulatory RNAs in Drosophila. Cell 130, 89–100 (2007).
Ruby, J. G., Jan, C. H. & Bartel, D. P. Intronic microRNA precursors that bypass Drosha processing. Nature 448, 83–86 (2007). Together with Okamura et al. (2007), this work reports the first non-canonical miRNA pathway, in which intron splicing bypasses Drosha to generate pre-miRNA mimics, termed mirtrons.
Bogerd, H. P. et al. A mammalian herpesvirus uses noncanonical expression and processing mechanisms to generate viral microRNAs. Mol. Cell 37, 135–142 (2010).
Xie, M. et al. Mammalian 5′-capped microRNA precursors that generate a single microRNA. Cell 155, 1568–1580 (2013).
Zamudio, J. R., Kelly, T. J. & Sharp, P. A. Argonaute-bound small RNAs from promoter-proximal RNA polymerase II. Cell 156, 920–934 (2014).
Cheloufi, S., Dos Santos, C. O., Chong, M. M. & Hannon, G. J. A dicer-independent miRNA biogenesis pathway that requires Ago catalysis. Nature 465, 584–589 (2010).
Cifuentes, D. et al. A novel miRNA processing pathway independent of dicer requires Argonaute2 catalytic activity. Science 328, 1694–1698 (2010).
Yang, J. S. et al. Conserved vertebrate mir-451 provides a platform for Dicer-independent, Ago2-mediated microRNA biogenesis. Proc. Natl Acad. Sci. USA 107, 15163–15168 (2010).
Acknowledgements
The authors thank G. La Rocca, B. Kleaveland and L. Joshua-Tor for critical reading, and the referees for informative comments. S.L. was supported by a training award from the NYSTEM contract #C32559GG and the Center for Stem Cell Biology at MSK. Work in E.C.L.’s group was supported by the National Institutes of Health (NIH) (R01-GM083300) and MSK Core Grant P30-CA008748. The authors apologize to those whose work is not included owing to space constraints.
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R.S., S.L., G.S. and E.C.L. wrote and edited the manuscript. R.S. S.L. and G.S. drafted the figures. R.S., S.L., G.S. and E.C.L. discussed and reviewed the manuscript before submission.
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Shang, R., Lee, S., Senavirathne, G. et al. microRNAs in action: biogenesis, function and regulation. Nat Rev Genet 24, 816–833 (2023). https://doi.org/10.1038/s41576-023-00611-y
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DOI: https://doi.org/10.1038/s41576-023-00611-y
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