- Email: [email protected]
Perspectives in targeting miRNA function
Bioorganic & Medicinal Chemistry 21 (2013) 6115–6118
Contents lists available at SciVerse ScienceDirect
Bioorganic & Medicinal Chemistry journal homepage: www.elsevier.com/locate/bmc
Perspectives in targeting miRNA function Christiane Schöniger, Christoph Arenz ⇑ Humboldt Universität zu Berlin, Institute for Chemistry, Brook-Taylor-Str. 2, 12489 Berlin, Germany
a r t i c l e
i n f o
Article history: Available online 2 April 2013 Keywords: miRNA Antagomirs Small molecules High throughput screening
a b s t r a c t First oligonucleotide analogues that inhibit miRNA function are currently investigated in clinical trials. In addition, several alternative methods are under development that may allow for controlling miRNA function by small molecules-mediated inhibiting of its biogenesis. In this perspectives article, we provide a short overview on recent developments in this field. Ó 2013 Elsevier Ltd. All rights reserved.
1. Introduction Micro-RNAs (miRNA) are an abundant class of endogenous, non-coding, regulatory RNA molecules of 21–23 nucleotides (nt) in length. Since their discovery in Caenohabditis elegans in 19931 and in humans in 2000,2 the interest in gaining control over miRNA-mediated regulation of gene expression has been steadily growing.3 Although the concept of targeting miRNAs as a potential therapy for human disease has already found its way into first clinical trials,4 many alternatives routes towards this end may exist and we obviously have just witnessed the very beginning of an emerging field in chemical biology with new modes of action and more classes of active molecules yet to come. It is believed that approximately 2000 human miRNAs exist, targeting around 30% of all expressed genes.5,6 Accordingly, miRNAs play a key role in cellular homeostasis and are involved in most if not all important cellular processes.7 Thus, it is not surprising that miRNA also has major impact on human health and disease. In fact, diverse cancers, cardiovascular or neuropsychiatric diseases like schizophrenia or depression correlate with altered miRNA expressions patterns.8–12 Under pathological conditions, the miRNA can either be down- or up-regulated and in a steadily growing number of reports, causative roles for miRNAs in disease development have clearly been shown.13,14 The obvious coherence of miRNA expression and human disease has prompted scientists to search for alternative RNA-directed therapeutic concepts, especially in diseases that have been connected to ‘undruggable’ enzymes or receptors. Up to now, several experimental strategies for manipulating miRNA function have been developed, which also include substitution with artificial miRNAs.15 Furthermore, a small molecule mediated enhancement of Dicing activity has been ⇑ Corresponding author. Tel.: +49 0 30 2093 8393; fax: +49 30 2093 6947. E-mail address: [email protected] (C. Arenz). 0968-0896/$ - see front matter Ó 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.bmc.2013.03.040
described,16 which unspecifically enhances RNAi as well as miRNA maturation. Herein, we will concentrate on the inhibition of miRNA function. From its initial transcription to its hybridization to target messenger RNAs (mRNAs), miRNAs interact with different proteins and enzymes in order to undergo a number of defined steps of maturation and translocation, all of which may be subjected to artificial chemical intervention. In this article, we summarize already established and more recent methods for miRNA targeting, with respect to the different steps of miRNA processing. 2. Biogenesis of miRNAs The biogenesis of miRNAs has been extensively reviewed.17 Briefly, its first step is usually the formation of a primary transcript by RNA polymerase II (Fig. 1)18 and the resulting primary miRNA (pri-miRNA) is then cleaved by the endonuclease Drosha to yield the about 70–80 nt long precursor miRNAs (pre-miRNA).19,20 These stem-loop shaped RNAs are subsequently translocated by Exportin 5 from the nucleus into the cytosol,21,22 where they are processed by the endonuclease Dicer to form the mature miRNAs.23 Upon Dicer cleavage, further proteins are recruited to form an RNA protein complex, which undergoes processing to the miRNA induced silencing complex (miRISC), harbouring only the miRNA’s active ‘guide’ strand, while the — usually not fully — complementary ‘passenger’ strand is subjected to degradation. The guide strand will find its target sequences within the 30 - untranslated regions (UTR) of mRNAs, which either leads to a translational arrest or alternatively will trigger the ‘Slicer’-activity of the miRISC causing mRNA degradation. 3. Mature miRNA as a target The historically first and also most obvious strategy to interfere with miRNA function is the direct targeting of mature miRNA. This approach, which already has found its way into clinical trials4
6116
C. Schöniger, C. Arenz / Bioorg. Med. Chem. 21 (2013) 6115–6118
Figure 1. Simplified overview over the miRNA biogenesis pathway and potential sites for inhibition (A–E) of miRNA activity: (A) miRNA transcription; (B) Drosha processing; (C) Dicer cleavage; (D) miRISC formation; (E) mRNA targeting.
makes use of anti-miRs or antagomirs. Antimir oligonucleotides (AMOs) are antisense-like oligonucleotide analogues of various types that are perfectly complementary to the mature miRNA guide strand.24,25 Upon hybridization with miRNA’s guide strand, these molecules have proved to reduce the effective concentration of miRNAs in a vast number of cellular models. The design of AMOs has been extensively reviewed.26 Special forms of AMOs are the chemically-modified antisense oligonucleotides like Antagomirs. The latter are 2-O-Me-RNAs conjugated to a single cholesterol moiety for better cellular uptake. Antagomirs have shown their therapeutic potential in one of the first in vivo experiments in mammals by effectively down-regulating cholesterol biosynthesis.27 Other nucleic acid analogues like the hydrophobic peptide nucleic acid (PNA) rely on conjugated polar amino acids like arginine or lysine, to ensure cellular uptake.28 The same rationale, but a completely different technology underlies a group of genetically-encoded repetitive miRNA binders, the so-called ‘microRNA sponges’, which have been introduced by the group of Sharp.29 These competitive miRNA binders, containing multiple tandem binding sites to a given miRNA, can be artificially expressed in cells, using vectors under the control of strong promoters, thereby specifically inhibiting miRNAs with a complementary heptameric seed. A single sponge can block an entire miRNA seed family, but with varying efficiencies, depending on their exact design. Alternatively, also single AMOs may be expressed from lentiviral vectors, which ensures the inhibition of miRNA function in cells for several days up to more than 1 month.30 Irrespective of the ongoing clinical trials, such approaches have revolutionized miRNA research by enabling easy and straight forward investigations in miRNA function in cell culture and in whole animals. An elegant addition to such techniques has been recently introduced by the Deiters group, who constructed photo-caged antagomirs, allowing for a light-inducible and spatio-temporal-controlled inhibition of miRNA function.31 4. Targeting miRNA maturation on level of Dicer cleavage Since the active mature miRNAs are only the final product, resulting from a number of consecutive enzyme-mediated steps, there may be alternatives to the direct targeting of miRNAs. An example is provided nature itself, where the endogenous protein lin-28 binds to pre-let-7, thereby inhibiting its maturation by Dicer in embryonic stem cells and embryocarcinoma cells.32 However,
the great hope that is connected with such approaches is the possibility of identifying small-molecules that may interfere with the maturation of specific miRNAs. A first example of a small molecule, protecting RNA from being subjected to Dicer cleavage was presented by the group of Yokobayashi. In their report, a small molecule specifically controlled Dicer-cleavage and the cellular activity of an aptamer-fused short hairpin RNA (shRNA) directed against a GFP reporter mRNA.33 Later, the group of Hartig similarly designed shRNAs with guanosine-rich (G-rich) sequences capable of folding into G-quadruplexes.34 Only some of the previously-described Gquadruplex binders like certain bis-quinolinium compounds or phorphyrazines were able to inhibit the cleavage of these shRNAs using recombinant Dicer, while being inactive within a cellular environment. In a first report focused on the inhibition of Dicermediated cleavage of pre-miRNA, our group described a homogenous assay for Dicer-mediated miRNA maturation, allowing for an easy in vitro screening of candidate molecules.35 This was achieved by Dicer-mediated cleavage of a doubly-labelled pre-miRNA, displaying a fluorophore on the 50 -end and a quencher moiety on 30 -end, which resulted in a Dicer dependent increase of fluorescence intensity. Using this assay, we exemplarily showed for the first time, that the aminoglycoside Kanamycin A at high concentrations was able to inhibit cleavage of pre-let7 from Drosophila melanogaster to the mature miRNA using recombinant Dicer.36 However, selectivity for a specific pre-miRNA was neither tested nor expected. Later, we synthesized six artificial Neamine- and 2Deoxystreptamine-dimers, which also were effective in this assay, with IC50-values in the sub-micromolar range.37 Again, specificity was not assessed. Due to the mechanism of Dicer, which recognizes the end of the pre-miRNA stem, the use of doubly-labelled premiRNAs as substrates harbours the risk of artificially-shifted Dicer cleavage sites, and the generation of screening artefacts. To meet this problem, we recently described a homogenous Dicer assay that relies on Dicer cleavage of unmodified pre-miRNAs and rolling circle amplification of the resulting mature miRNAs.38 Interestingly, the inhibitory potency of the molecules described above was essentially the same, independently of which assay was used. In a different study by the group of Herdewijn, the effect of several types of molecules on Dicer-mediated cleavage of a radioactively labelled pre-miRNA was tested.39 Furthermore, in a very recent report the potential of various Xanthone derivatives as potential inhibitors of miRNA processing by human Dicer was assessed by
C. Schöniger, C. Arenz / Bioorg. Med. Chem. 21 (2013) 6115–6118
the group of Nakatani.40 A peak in this story was reached with the recent report that the aminoglycoside Streptomycin selectively inhibits Dicer-mediated maturation of miR-21 in living cells.41 This finding was the result of a top-down approach, starting with a primary screening of commercially available aminoglycosides for modulation of miR-21-controlled luciferase activity as previously applied by Deiters and co-workers.42 In a second step, specificity and mechanism were confirmed by inhibition of pre-miR-21 cleavage by recombinant Dicer in a radioactive assay format. Specificity of binding to pre-miR-21 was also confirmed by means of footprinting assays and by molecular modelling, respectively. Further molecules holding great promise for an inhibition of Dicer-mediated pre-miRNA cleavage have been described, but instead of assaying miRNA maturation, only the binding to premiRNAs and related hairpin RNAs was described: The group of Beal synthesized macrocyclic helix threading peptides (HTP) that target certain duplex RNA structures selectively by threading intercalation.43 Similarly, Luebke et. al. identified specific ligands to the hairpin of pre-miR-21 after screening a library of more than 7500 N-substituted oligoglycines (peptoids), but their potential as inhibitors of miR-21 maturation was not evaluated.44 Aside from these small-molecule approaches, an inhibition of miRNA maturation may also be obtained by oligonucleotides. The first example was presented by Plasterk and coworkers who designed long morpholino oligonucleotide analogues not only targeting the mature miR-375, but also the Dicer and Drosha cleavage sites in the respective miRNA precursor molecules to obtain an inhibition of miRNA maturation in living zebrafish.45 A similar approach was recently reported by the group of Romanelli, who conjugated PNAs targeting pre-miRNAs with peptide sequences for effective cellular delivery.46 Not only was miRNA activity inhibited, but also concentration of mature miRNAs was effectively reduced. A completely different approach for targeting pre-miRNAs was developed by Maiti and co-workers, who engineered oligonucleotide enzymes (DNAzymes and LNAzymes) recognizing and cleaving different human pre-miRNAs. These so-called antagomirzymes led to a reduction of miRNA activity in living cells, but only at very high concentrations.47 5. Targeting miRNA maturation on level of Drosha cleavage As already mentioned above, morpholino oligonuclotides have been shown to inhibit Drosha mediated cleavage of pri-miRNAs in zebrafish.45 In an independent approach, a group of 20 -O-methyl oligoribonucleotides complementary to conserved apical loops of pri-miRNAs (looptomiRs) was designed, inhibiting pri-miRNA processing in vitro using nuclear extracts from HeLa cells.48 Similarly, RNA aptamers targeting the apical-loop domain of pri-miRNA18a were isolated and characterized by the groups of Mayer and Famulok.49 The aptamers not only bound to pri-miR-18a, but also inhibited the biogenesis of the pre-miRNA in HeLa nuclear extracts, while the mechanism appeared to be distinct from that of the previously described looptomiRs. For both types of molecules, no function in living cells was reported. 6. Miscellaneous approaches towards an inhibition of miRNA function The first small molecules inhibiting the activity of specific miRNAs in living cells have been described by Deiters and co-workers.42 They constructed a reporter gene vector, resulting in a luciferase mRNA harbouring miR-21 binding sites in its 30 UTR. Inhibition of miR-21 function would produce fully active luciferase, while under control conditions luciferase expression remained silenced. Screening of a commercial small-molecule library
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of >1200 compounds yielded in an active diazobenzene structural motif, which led to a small molecule compound selectively inhibiting the function of miR-21 in the low micromolar range. A similar screening of 1364 compounds also yielded an inhibitor of the liver-specific miR-122.50 In both cases however, the exact molecular targets, which are obviously upstream of the pri-miRNA synthesis, remain unclear. The great advantage of this strategy is the possibility to identify effective modulators of miRNA function in high-throughput screenings (HTS).51 Recently, a similar endeavour was reported using GFP as a reporter gene, allowing for imageguided high content screens at very low hit-rates.52 In another recent study, more than 1200 compounds were screened for an inhibition of RISC loading. Interestingly three molecules affected assembly of the RISC or miRISC at sub-micromolar concentrations in vitro and in living cells thereby enabling mechanistic studies or global inhibition of miRNA activity.53 7. Conclusion While inhibition of miRNA activity is already subject to clinical trials, the number of alternative small-molecule-based methods to interfere with miRNA biogenesis in vitro and in cultured cells is steadily increasing. In most cases however, more specific and more potent molecules will be needed in order to provide alternatives to modulation of miRNA activity by means of oligonucleotide analogues. References and notes 1. Lee, R. C.; Feinbaum, R. L.; Ambros, V. Cell 1993, 75, 843. 2. Pasquinelli, A. E.; Reinhart, B. J.; Slack, F.; Martindale, M. Q.; Kuroda, M. I.; Maller, B.; Hayward, D. C.; Ball, E. E.; Degnan, B.; Muller, P.; Spring, J.; Srinivasan, A.; Fishman, M.; Finnerty, J.; Corbo, J.; Levine, M.; Leahy, P.; Davidson, E.; Ruvkun, G. Nature 2000, 408, 86. 3. Arenz, C. Angew. Chem., Int. Ed. 2006, 45, 5048. 4. Schoof, C. R.; Botelho, E. L.; Izzotti, A.; Vasques Ldos, R. Am. J. Cancer Res. 2012, 2, 414. 5. Bentwich, I.; Avniel, A.; Karov, Y.; Aharonov, R.; Gilad, S.; Barad, O.; Barzilai, A.; Einat, P.; Einav, U.; Meiri, E.; Sharon, E.; Spector, Y.; Bentwich, Z. Nat. Genet. 2005, 37, 766. 6. Lewis, B. P.; Burge, C. B.; Bartel, D. P. Cell 2005, 120, 15. 7. He, L.; Hannon, G. J. Nat. Rev. Genet. 2004, 5, 522. 8. Croce, C. M.; Calin, G. A. Cell 2005, 122, 6. 9. Bicker, S.; Schratt, G. J. Cell. Mol. Med. 2008, 12, 1466. 10. Fiore, R.; Siegel, G.; Schratt, G. Biochim. Biophys. Acta 2008, 1779, 471. 11. Thum, T.; Gross, C.; Fiedler, J.; Fischer, T.; Kissler, S.; Bussen, M.; Galuppo, P.; Just, S.; Rottbauer, W.; Frantz, S.; Castoldi, M.; Soutschek, J.; Koteliansky, V.; Rosenwald, A.; Basson, M. A.; Licht, J. D.; Pena, J. T. R.; Rouhanifard, S. H.; Muckenthaler, M. U.; Tuschl, T.; Martin, G. R.; Bauersachs, J.; Engelhardt, S. Nature 2008, 456, 980. 12. Wang, C.; Bian, Z.; Wei, D.; Zhang, J.-g. Mol. Cell. Biochem. 2011, 352, 197. 13. Asangani, I. A.; Rasheed, S. A. K.; Nikolova, D. A.; Leupold, J. H.; Colburn, N. H.; Post, S.; Allgayer, H. Oncogene 2007, 27, 2128. 14. O’Connell, R. M.; Rao, D. S.; Chaudhuri, A. A.; Boldin, M. P.; Taganov, K. D.; Nicoll, J.; Paquette, R. L.; Baltimore, D. J. Exp. Med. 2008, 205, 585. 15. Trang, P.; Wiggins, J. F.; Daige, C. L.; Cho, C.; Omotola, M.; Brown, D.; Weidhaas, J. B.; Bader, A. G.; Slack, F. J. Mol. Ther. 2011, 19, 1116. 16. Shan, G.; Li, Y.; Zhang, J.; Li, W.; Szulwach, K. E.; Duan, R.; Faghihi, M. A.; Khalil, A. M.; Lu, L.; Paroo, Z.; Chan, A. W.; Shi, Z.; Liu, Q.; Wahlestedt, C.; He, C.; Jin, P. Nat. Biotechnol. 2008, 26, 933. 17. Winter, J.; Jung, S.; Keller, S.; Gregory, R. I.; Diederichs, S. Nat. Cell Biol. 2009, 11, 228. 18. Lee, Y.; Kim, M.; Han, J.; Yeom, K.-H.; Lee, S.; Baek, S. H.; Kim, V. N. EMBO J. 2004, 23, 4051. 19. Han, J.; Lee, Y.; Yeom, K. H.; Kim, Y. K.; Jin, H.; Kim, V. N. Genes Dev. 2004, 18, 3016. 20. Lee, Y.; Ahn, C.; Han, J.; Choi, H.; Kim, J.; Yim, J.; Lee, J.; Provost, P.; Radmark, O.; Kim, S.; Kim, V. N. Nature 2003, 425, 415. 21. Yi, R.; Qin, Y.; Macara, I. G.; Cullen, B. R. Genes Dev. 2003, 17, 3011. 22. Lund, E.; Guttinger, S.; Calado, A.; Dahlberg, J. E.; Kutay, U. Science 2004, 303, 95. 23. Bernstein, E.; Caudy, A. A.; Hammond, S. M.; Hannon, G. J. Nature 2001, 409, 363. 24. Meister, G.; Landthaler, M.; Dorsett, Y.; Tuschl, T. RNA 2004, 10, 544. 25. Weiler, J.; Hunziker, J.; Hall, J. Gene Ther. 2005, 13, 496. 26. Lennox, K. A.; Behlke, M. A. Gene Ther. 2011, 18, 1111. 27. Krutzfeldt, J.; Rajewsky, N.; Braich, R.; Rajeev, K. G.; Tuschl, T.; Manoharan, M.; Stoffel, M. Nature 2005, 438, 685.
6118 28. 29. 30. 31. 32. 33. 34. 35. 36. 37. 38. 39. 40. 41.
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Torres, A. G.; Threlfall, R. N.; Gait, M. J. Artif DNA PNA XNA 2011, 2, 71. Ebert, M. S.; Neilson, J. R.; Sharp, P. A. Nat. Methods 2007, 4, 721. Haraguchi, T.; Ozaki, Y.; Iba, H. Nucleic Acids Res. 2009, 37, e43. Connelly, C. M.; Uprety, R.; Hemphill, J.; Deiters, A. Mol. Biosyst. 2012, 8, 2987. Rybak, A.; Fuchs, H.; Smirnova, L.; Brandt, C.; Pohl, E. E.; Nitsch, R.; Wulczyn, F. G. Nat. Cell Biol. 2008, 10, 987. An, C. I.; Trinh, V. B.; Yokobayashi, Y. RNA 2006, 12, 710. Henn, A.; Joachimi, A.; Goncalves, D. P.; Monchaud, D.; Teulade-Fichou, M. P.; Sanders, J. K.; Hartig, J. S. Chembiochem 2008, 9, 2722. Davies, B. P.; Arenz, C. Angew. Chem., Int. Ed. 2006, 45, 5550. Davies, B. P.; Arenz, C. Bioorg. Med. Chem. 2008, 16, 49. Klemm, C. M.; Berthelmann, A.; Neubacher, S.; Arenz, C. Eur. J. Org. Chem. 2009, 2788. Neubacher, S.; Dojahn, C. M.; Arenz, C. ChemBioChem 2011, 12, 2302. Maiti, M.; Nauwelaerts, K.; Herdewijn, P. Bioorg. Med. Chem. Lett. 2012, 22, 1709. Murata, A.; Fukuzumi, T.; Umemoto, S.; Nakatani, K. Bioorg. Med. Chem. Lett. 2013, 23, 252. Bose, D.; Jayaraj, G.; Suryawanshi, H.; Agarwala, P.; Pore, S. K.; Banerjee, R.; Maiti, S. Angew. Chem., Int. Ed. 2012, 51, 1019.
42. Gumireddy, K.; Young, D. D.; Xiong, X.; Hogenesch, J. B.; Huang, Q.; Deiters, A. Angew. Chem., Int. Ed. 2008, 47, 7482. 43. Krishnamurthy, M.; Simon, K.; Orendt, A. M.; Beal, P. A. Angew. Chem., Int. Ed. 2007, 46, 7044. 44. Chirayil, S.; Chirayil, R.; Luebke, K. J. Nucleic Acids Res. 2009, 37, 5486. 45. Kloosterman, W. P.; Lagendijk, A. K.; Ketting, R. F.; Moulton, J. D.; Plasterk, R. H. PLoS Biol. 2007, 5, e203. 46. Jadhav, V. M.; Scaria, V.; Maiti, S. Angew. Chem., Int. Ed. 2009, 48, 2557. 47. Avitabile, C.; Saviano, M.; D’Andrea, L.; Bianchi, N.; Fabbri, E.; Brognara, E.; Gambari, R.; Romanelli, A. Artif DNA PNA XNA 2012, 3, 88. 48. Michlewski, G.; Guil, S.; Semple, C. A.; Caceres, J. F. Mol. Cell 2008, 32, 383. 49. Lunse, C. E.; Michlewski, G.; Hopp, C. S.; Rentmeister, A.; Caceres, J. F.; Famulok, M.; Mayer, G. Angew. Chem., Int. Ed. 2010, 49, 4674. 50. Young, D. D.; Connelly, C. M.; Grohmann, C.; Deiters, A. J. Am. Chem. Soc. 2010, 132, 7976. 51. Connelly, C. M.; Thomas, M.; Deiters, A. J. Biomol. Screen. 2012, 17, 822. 52. Shum, D.; Bhinder, B.; Radu, C.; Calder, P.; Ramirez, C. N.; Djaballah, H. Comb. Chem. High Throughput Screen. 2012, 15, 529. 53. Tan, G. S.; Chiu, C. H.; Garchow, B. G.; Metzler, D.; Diamond, S. L.; Kiriakidou, M. ACS Chem. Biol. 2012, 7, 403.
Contents lists available at SciVerse ScienceDirect
Bioorganic & Medicinal Chemistry journal homepage: www.elsevier.com/locate/bmc
Perspectives in targeting miRNA function Christiane Schöniger, Christoph Arenz ⇑ Humboldt Universität zu Berlin, Institute for Chemistry, Brook-Taylor-Str. 2, 12489 Berlin, Germany
a r t i c l e
i n f o
Article history: Available online 2 April 2013 Keywords: miRNA Antagomirs Small molecules High throughput screening
a b s t r a c t First oligonucleotide analogues that inhibit miRNA function are currently investigated in clinical trials. In addition, several alternative methods are under development that may allow for controlling miRNA function by small molecules-mediated inhibiting of its biogenesis. In this perspectives article, we provide a short overview on recent developments in this field. Ó 2013 Elsevier Ltd. All rights reserved.
1. Introduction Micro-RNAs (miRNA) are an abundant class of endogenous, non-coding, regulatory RNA molecules of 21–23 nucleotides (nt) in length. Since their discovery in Caenohabditis elegans in 19931 and in humans in 2000,2 the interest in gaining control over miRNA-mediated regulation of gene expression has been steadily growing.3 Although the concept of targeting miRNAs as a potential therapy for human disease has already found its way into first clinical trials,4 many alternatives routes towards this end may exist and we obviously have just witnessed the very beginning of an emerging field in chemical biology with new modes of action and more classes of active molecules yet to come. It is believed that approximately 2000 human miRNAs exist, targeting around 30% of all expressed genes.5,6 Accordingly, miRNAs play a key role in cellular homeostasis and are involved in most if not all important cellular processes.7 Thus, it is not surprising that miRNA also has major impact on human health and disease. In fact, diverse cancers, cardiovascular or neuropsychiatric diseases like schizophrenia or depression correlate with altered miRNA expressions patterns.8–12 Under pathological conditions, the miRNA can either be down- or up-regulated and in a steadily growing number of reports, causative roles for miRNAs in disease development have clearly been shown.13,14 The obvious coherence of miRNA expression and human disease has prompted scientists to search for alternative RNA-directed therapeutic concepts, especially in diseases that have been connected to ‘undruggable’ enzymes or receptors. Up to now, several experimental strategies for manipulating miRNA function have been developed, which also include substitution with artificial miRNAs.15 Furthermore, a small molecule mediated enhancement of Dicing activity has been ⇑ Corresponding author. Tel.: +49 0 30 2093 8393; fax: +49 30 2093 6947. E-mail address: [email protected] (C. Arenz). 0968-0896/$ - see front matter Ó 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.bmc.2013.03.040
described,16 which unspecifically enhances RNAi as well as miRNA maturation. Herein, we will concentrate on the inhibition of miRNA function. From its initial transcription to its hybridization to target messenger RNAs (mRNAs), miRNAs interact with different proteins and enzymes in order to undergo a number of defined steps of maturation and translocation, all of which may be subjected to artificial chemical intervention. In this article, we summarize already established and more recent methods for miRNA targeting, with respect to the different steps of miRNA processing. 2. Biogenesis of miRNAs The biogenesis of miRNAs has been extensively reviewed.17 Briefly, its first step is usually the formation of a primary transcript by RNA polymerase II (Fig. 1)18 and the resulting primary miRNA (pri-miRNA) is then cleaved by the endonuclease Drosha to yield the about 70–80 nt long precursor miRNAs (pre-miRNA).19,20 These stem-loop shaped RNAs are subsequently translocated by Exportin 5 from the nucleus into the cytosol,21,22 where they are processed by the endonuclease Dicer to form the mature miRNAs.23 Upon Dicer cleavage, further proteins are recruited to form an RNA protein complex, which undergoes processing to the miRNA induced silencing complex (miRISC), harbouring only the miRNA’s active ‘guide’ strand, while the — usually not fully — complementary ‘passenger’ strand is subjected to degradation. The guide strand will find its target sequences within the 30 - untranslated regions (UTR) of mRNAs, which either leads to a translational arrest or alternatively will trigger the ‘Slicer’-activity of the miRISC causing mRNA degradation. 3. Mature miRNA as a target The historically first and also most obvious strategy to interfere with miRNA function is the direct targeting of mature miRNA. This approach, which already has found its way into clinical trials4
6116
C. Schöniger, C. Arenz / Bioorg. Med. Chem. 21 (2013) 6115–6118
Figure 1. Simplified overview over the miRNA biogenesis pathway and potential sites for inhibition (A–E) of miRNA activity: (A) miRNA transcription; (B) Drosha processing; (C) Dicer cleavage; (D) miRISC formation; (E) mRNA targeting.
makes use of anti-miRs or antagomirs. Antimir oligonucleotides (AMOs) are antisense-like oligonucleotide analogues of various types that are perfectly complementary to the mature miRNA guide strand.24,25 Upon hybridization with miRNA’s guide strand, these molecules have proved to reduce the effective concentration of miRNAs in a vast number of cellular models. The design of AMOs has been extensively reviewed.26 Special forms of AMOs are the chemically-modified antisense oligonucleotides like Antagomirs. The latter are 2-O-Me-RNAs conjugated to a single cholesterol moiety for better cellular uptake. Antagomirs have shown their therapeutic potential in one of the first in vivo experiments in mammals by effectively down-regulating cholesterol biosynthesis.27 Other nucleic acid analogues like the hydrophobic peptide nucleic acid (PNA) rely on conjugated polar amino acids like arginine or lysine, to ensure cellular uptake.28 The same rationale, but a completely different technology underlies a group of genetically-encoded repetitive miRNA binders, the so-called ‘microRNA sponges’, which have been introduced by the group of Sharp.29 These competitive miRNA binders, containing multiple tandem binding sites to a given miRNA, can be artificially expressed in cells, using vectors under the control of strong promoters, thereby specifically inhibiting miRNAs with a complementary heptameric seed. A single sponge can block an entire miRNA seed family, but with varying efficiencies, depending on their exact design. Alternatively, also single AMOs may be expressed from lentiviral vectors, which ensures the inhibition of miRNA function in cells for several days up to more than 1 month.30 Irrespective of the ongoing clinical trials, such approaches have revolutionized miRNA research by enabling easy and straight forward investigations in miRNA function in cell culture and in whole animals. An elegant addition to such techniques has been recently introduced by the Deiters group, who constructed photo-caged antagomirs, allowing for a light-inducible and spatio-temporal-controlled inhibition of miRNA function.31 4. Targeting miRNA maturation on level of Dicer cleavage Since the active mature miRNAs are only the final product, resulting from a number of consecutive enzyme-mediated steps, there may be alternatives to the direct targeting of miRNAs. An example is provided nature itself, where the endogenous protein lin-28 binds to pre-let-7, thereby inhibiting its maturation by Dicer in embryonic stem cells and embryocarcinoma cells.32 However,
the great hope that is connected with such approaches is the possibility of identifying small-molecules that may interfere with the maturation of specific miRNAs. A first example of a small molecule, protecting RNA from being subjected to Dicer cleavage was presented by the group of Yokobayashi. In their report, a small molecule specifically controlled Dicer-cleavage and the cellular activity of an aptamer-fused short hairpin RNA (shRNA) directed against a GFP reporter mRNA.33 Later, the group of Hartig similarly designed shRNAs with guanosine-rich (G-rich) sequences capable of folding into G-quadruplexes.34 Only some of the previously-described Gquadruplex binders like certain bis-quinolinium compounds or phorphyrazines were able to inhibit the cleavage of these shRNAs using recombinant Dicer, while being inactive within a cellular environment. In a first report focused on the inhibition of Dicermediated cleavage of pre-miRNA, our group described a homogenous assay for Dicer-mediated miRNA maturation, allowing for an easy in vitro screening of candidate molecules.35 This was achieved by Dicer-mediated cleavage of a doubly-labelled pre-miRNA, displaying a fluorophore on the 50 -end and a quencher moiety on 30 -end, which resulted in a Dicer dependent increase of fluorescence intensity. Using this assay, we exemplarily showed for the first time, that the aminoglycoside Kanamycin A at high concentrations was able to inhibit cleavage of pre-let7 from Drosophila melanogaster to the mature miRNA using recombinant Dicer.36 However, selectivity for a specific pre-miRNA was neither tested nor expected. Later, we synthesized six artificial Neamine- and 2Deoxystreptamine-dimers, which also were effective in this assay, with IC50-values in the sub-micromolar range.37 Again, specificity was not assessed. Due to the mechanism of Dicer, which recognizes the end of the pre-miRNA stem, the use of doubly-labelled premiRNAs as substrates harbours the risk of artificially-shifted Dicer cleavage sites, and the generation of screening artefacts. To meet this problem, we recently described a homogenous Dicer assay that relies on Dicer cleavage of unmodified pre-miRNAs and rolling circle amplification of the resulting mature miRNAs.38 Interestingly, the inhibitory potency of the molecules described above was essentially the same, independently of which assay was used. In a different study by the group of Herdewijn, the effect of several types of molecules on Dicer-mediated cleavage of a radioactively labelled pre-miRNA was tested.39 Furthermore, in a very recent report the potential of various Xanthone derivatives as potential inhibitors of miRNA processing by human Dicer was assessed by
C. Schöniger, C. Arenz / Bioorg. Med. Chem. 21 (2013) 6115–6118
the group of Nakatani.40 A peak in this story was reached with the recent report that the aminoglycoside Streptomycin selectively inhibits Dicer-mediated maturation of miR-21 in living cells.41 This finding was the result of a top-down approach, starting with a primary screening of commercially available aminoglycosides for modulation of miR-21-controlled luciferase activity as previously applied by Deiters and co-workers.42 In a second step, specificity and mechanism were confirmed by inhibition of pre-miR-21 cleavage by recombinant Dicer in a radioactive assay format. Specificity of binding to pre-miR-21 was also confirmed by means of footprinting assays and by molecular modelling, respectively. Further molecules holding great promise for an inhibition of Dicer-mediated pre-miRNA cleavage have been described, but instead of assaying miRNA maturation, only the binding to premiRNAs and related hairpin RNAs was described: The group of Beal synthesized macrocyclic helix threading peptides (HTP) that target certain duplex RNA structures selectively by threading intercalation.43 Similarly, Luebke et. al. identified specific ligands to the hairpin of pre-miR-21 after screening a library of more than 7500 N-substituted oligoglycines (peptoids), but their potential as inhibitors of miR-21 maturation was not evaluated.44 Aside from these small-molecule approaches, an inhibition of miRNA maturation may also be obtained by oligonucleotides. The first example was presented by Plasterk and coworkers who designed long morpholino oligonucleotide analogues not only targeting the mature miR-375, but also the Dicer and Drosha cleavage sites in the respective miRNA precursor molecules to obtain an inhibition of miRNA maturation in living zebrafish.45 A similar approach was recently reported by the group of Romanelli, who conjugated PNAs targeting pre-miRNAs with peptide sequences for effective cellular delivery.46 Not only was miRNA activity inhibited, but also concentration of mature miRNAs was effectively reduced. A completely different approach for targeting pre-miRNAs was developed by Maiti and co-workers, who engineered oligonucleotide enzymes (DNAzymes and LNAzymes) recognizing and cleaving different human pre-miRNAs. These so-called antagomirzymes led to a reduction of miRNA activity in living cells, but only at very high concentrations.47 5. Targeting miRNA maturation on level of Drosha cleavage As already mentioned above, morpholino oligonuclotides have been shown to inhibit Drosha mediated cleavage of pri-miRNAs in zebrafish.45 In an independent approach, a group of 20 -O-methyl oligoribonucleotides complementary to conserved apical loops of pri-miRNAs (looptomiRs) was designed, inhibiting pri-miRNA processing in vitro using nuclear extracts from HeLa cells.48 Similarly, RNA aptamers targeting the apical-loop domain of pri-miRNA18a were isolated and characterized by the groups of Mayer and Famulok.49 The aptamers not only bound to pri-miR-18a, but also inhibited the biogenesis of the pre-miRNA in HeLa nuclear extracts, while the mechanism appeared to be distinct from that of the previously described looptomiRs. For both types of molecules, no function in living cells was reported. 6. Miscellaneous approaches towards an inhibition of miRNA function The first small molecules inhibiting the activity of specific miRNAs in living cells have been described by Deiters and co-workers.42 They constructed a reporter gene vector, resulting in a luciferase mRNA harbouring miR-21 binding sites in its 30 UTR. Inhibition of miR-21 function would produce fully active luciferase, while under control conditions luciferase expression remained silenced. Screening of a commercial small-molecule library
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of >1200 compounds yielded in an active diazobenzene structural motif, which led to a small molecule compound selectively inhibiting the function of miR-21 in the low micromolar range. A similar screening of 1364 compounds also yielded an inhibitor of the liver-specific miR-122.50 In both cases however, the exact molecular targets, which are obviously upstream of the pri-miRNA synthesis, remain unclear. The great advantage of this strategy is the possibility to identify effective modulators of miRNA function in high-throughput screenings (HTS).51 Recently, a similar endeavour was reported using GFP as a reporter gene, allowing for imageguided high content screens at very low hit-rates.52 In another recent study, more than 1200 compounds were screened for an inhibition of RISC loading. Interestingly three molecules affected assembly of the RISC or miRISC at sub-micromolar concentrations in vitro and in living cells thereby enabling mechanistic studies or global inhibition of miRNA activity.53 7. Conclusion While inhibition of miRNA activity is already subject to clinical trials, the number of alternative small-molecule-based methods to interfere with miRNA biogenesis in vitro and in cultured cells is steadily increasing. In most cases however, more specific and more potent molecules will be needed in order to provide alternatives to modulation of miRNA activity by means of oligonucleotide analogues. References and notes 1. Lee, R. C.; Feinbaum, R. L.; Ambros, V. Cell 1993, 75, 843. 2. Pasquinelli, A. E.; Reinhart, B. J.; Slack, F.; Martindale, M. Q.; Kuroda, M. I.; Maller, B.; Hayward, D. C.; Ball, E. E.; Degnan, B.; Muller, P.; Spring, J.; Srinivasan, A.; Fishman, M.; Finnerty, J.; Corbo, J.; Levine, M.; Leahy, P.; Davidson, E.; Ruvkun, G. Nature 2000, 408, 86. 3. Arenz, C. Angew. Chem., Int. Ed. 2006, 45, 5048. 4. Schoof, C. R.; Botelho, E. L.; Izzotti, A.; Vasques Ldos, R. Am. J. Cancer Res. 2012, 2, 414. 5. Bentwich, I.; Avniel, A.; Karov, Y.; Aharonov, R.; Gilad, S.; Barad, O.; Barzilai, A.; Einat, P.; Einav, U.; Meiri, E.; Sharon, E.; Spector, Y.; Bentwich, Z. Nat. Genet. 2005, 37, 766. 6. Lewis, B. P.; Burge, C. B.; Bartel, D. P. Cell 2005, 120, 15. 7. He, L.; Hannon, G. J. Nat. Rev. Genet. 2004, 5, 522. 8. Croce, C. M.; Calin, G. A. Cell 2005, 122, 6. 9. Bicker, S.; Schratt, G. J. Cell. Mol. Med. 2008, 12, 1466. 10. Fiore, R.; Siegel, G.; Schratt, G. Biochim. Biophys. Acta 2008, 1779, 471. 11. Thum, T.; Gross, C.; Fiedler, J.; Fischer, T.; Kissler, S.; Bussen, M.; Galuppo, P.; Just, S.; Rottbauer, W.; Frantz, S.; Castoldi, M.; Soutschek, J.; Koteliansky, V.; Rosenwald, A.; Basson, M. A.; Licht, J. D.; Pena, J. T. R.; Rouhanifard, S. H.; Muckenthaler, M. U.; Tuschl, T.; Martin, G. R.; Bauersachs, J.; Engelhardt, S. Nature 2008, 456, 980. 12. Wang, C.; Bian, Z.; Wei, D.; Zhang, J.-g. Mol. Cell. Biochem. 2011, 352, 197. 13. Asangani, I. A.; Rasheed, S. A. K.; Nikolova, D. A.; Leupold, J. H.; Colburn, N. H.; Post, S.; Allgayer, H. Oncogene 2007, 27, 2128. 14. O’Connell, R. M.; Rao, D. S.; Chaudhuri, A. A.; Boldin, M. P.; Taganov, K. D.; Nicoll, J.; Paquette, R. L.; Baltimore, D. J. Exp. Med. 2008, 205, 585. 15. Trang, P.; Wiggins, J. F.; Daige, C. L.; Cho, C.; Omotola, M.; Brown, D.; Weidhaas, J. B.; Bader, A. G.; Slack, F. J. Mol. Ther. 2011, 19, 1116. 16. Shan, G.; Li, Y.; Zhang, J.; Li, W.; Szulwach, K. E.; Duan, R.; Faghihi, M. A.; Khalil, A. M.; Lu, L.; Paroo, Z.; Chan, A. W.; Shi, Z.; Liu, Q.; Wahlestedt, C.; He, C.; Jin, P. Nat. Biotechnol. 2008, 26, 933. 17. Winter, J.; Jung, S.; Keller, S.; Gregory, R. I.; Diederichs, S. Nat. Cell Biol. 2009, 11, 228. 18. Lee, Y.; Kim, M.; Han, J.; Yeom, K.-H.; Lee, S.; Baek, S. H.; Kim, V. N. EMBO J. 2004, 23, 4051. 19. Han, J.; Lee, Y.; Yeom, K. H.; Kim, Y. K.; Jin, H.; Kim, V. N. Genes Dev. 2004, 18, 3016. 20. Lee, Y.; Ahn, C.; Han, J.; Choi, H.; Kim, J.; Yim, J.; Lee, J.; Provost, P.; Radmark, O.; Kim, S.; Kim, V. N. Nature 2003, 425, 415. 21. Yi, R.; Qin, Y.; Macara, I. G.; Cullen, B. R. Genes Dev. 2003, 17, 3011. 22. Lund, E.; Guttinger, S.; Calado, A.; Dahlberg, J. E.; Kutay, U. Science 2004, 303, 95. 23. Bernstein, E.; Caudy, A. A.; Hammond, S. M.; Hannon, G. J. Nature 2001, 409, 363. 24. Meister, G.; Landthaler, M.; Dorsett, Y.; Tuschl, T. RNA 2004, 10, 544. 25. Weiler, J.; Hunziker, J.; Hall, J. Gene Ther. 2005, 13, 496. 26. Lennox, K. A.; Behlke, M. A. Gene Ther. 2011, 18, 1111. 27. Krutzfeldt, J.; Rajewsky, N.; Braich, R.; Rajeev, K. G.; Tuschl, T.; Manoharan, M.; Stoffel, M. Nature 2005, 438, 685.
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