The test system utilizes fluorophore-labeled substrate RNAs to analyze the processing reaction by automated, high-throughput capillary electrophoresis (CE) in combination with a laser-induced fluorescence (LIF) readout. We optimized the assay for high-throughput screening (HTS)-experiments and devised a multiplex fluorophore-labeling regime to scrutinize the U-insertion/U-deletion reaction simultaneously. The assay is robust, it requires only nanogram amounts of materials and it meets all performance criteria for HTS-methods. As such the test system should be helpful in the search for trypanosome-specific pharmaceuticals.Mitochondrial gene expression in African trypanosomes and other trypanosomatid pathogens requires a U-nucleotide specific insertion/deletion-type RNA-editing reaction. The process is catalyzed by a macromolecular protein complex known as the editosome. Editosomes are restricted to the trypanosomatid clade and since editing is essential for the parasites, the protein complex represents a near perfect target for drug intervention strategies. Here, we report the development of an improved  Trypanosoma brucei, an extracellular, single-cell parasite. The organism proliferates in the blood and lymph fluid and is capable of crossing the blood-brain barrier where it induces a wide spectrum of neurological symptoms (Human African trypanosomiasis (HAT), also known as sleeping sickness, is a neglected tropical disease. Despite a decreasing number of new infections in recent years, still 70 million people in 36 African countries are at risk of becoming infected . In respsymptoms . Left unsymptoms .Mitochondrial gene expression in trypanosomes and related pathogens requires an RNA-editing reaction in which sequence-deficient, non-translatable primary transcripts are converted into translatable mRNAs by the site-specific insertion and deletion of exclusively U-nucleotides . The reain vitro RNA-editing assays dimethylsilyl (TBDMS)-protected phosphoramidites. 5-Carboxytetramethyl-rhodamine , 6-hexachloro-fluorescein  and 6-carboxy-fluorescein  were chosen as fluorophore substituents and were introduced post-synthetically either at the 5′- or 3′-ends of the different oligoribonucleotides. For that the RNA-oligos were synthesized to contain either a 5′- or 3′-terminal C6-aminolinker. The primary amino group was then conjugated to carboxyl functional groups of FAM, TAMRA and HEX using an EDC (1-ethyl-3-(3-dimethylaminopropyl)carbodiimide)-mediated coupling reaction. As additional modifications, some RNAs were synthesized to include up to six phosphorothioate (PS) backbone modifications to increase their RNase-stability. Similarly, some of the 3′-pre-mRNA oligonucleotides were 3′-end modified by introducing a hexamethylene-amino-linker. Base-protecting groups were removed at mild conditions using NH4OH/EtOH (3:1) at RT and 2′-silyl protecting groups were removed using neat triethylamine trihydrofluoride. All RNA-oligonucleotides were HPLC-purified, analyzed by mass-spectrometry and further scrutinized in denaturing polyacrylamide gels (A260) using the molar extinction coefficients (ϵ in l/mol cm) listed below. Oligoribonucleotides representing 3′-pre-mRNA fragments were enzymatically 5′-phosphorylated using T4-polynucleotide kinase (T4-PNK) and ATP using standard conditions. The following RNA-oligonucleotides were synthesized:RNA-oligonucleotides were synthesized by solid-phase synthesis on controlled pore glass (CPG)-beads  using 2′ide gels . RNA-con2EDTA (TE) pH 7.5 followed by denaturation at 75°C for 2 min. Denatured oligoribonucleotides were annealed by cooling samples to 27°C at a rate of 0.08°C/s. Formation of the pre-mRNA/gRNA-hybrid RNAs was verified by electrophoresis in native 15% (w/v) polyacrylamide gels followed by staining with Toluidine blue O. Stained gels were densitometrically analyzed.Trimolecular pre-mRNA/gRNA-hybrid molecules were generated by hybridization of the individual 5′- and 3′-pre-mRNA oligoribonucleotides to base complementary gRNA molecules. For that, equimolar amounts (4 pmol) of the three RNA-oligonucleotides were combined in a final volume of 0.1 ml 10 mM Tris/HCl, 1mM NaTrypanosoma brucei), which were grown at 27°C in SDM-79 medium  as described in .Editosomes were purified from insect-stage African trypanosomes  supplemented with 0.5 mM DTT and 0.5 mM ATP. For the U-insertion assay, additional 0.1 mM UTP or analogs of UTP were added. Reactions contained 0.4 pmol pre-annealed pre-mRNA/gRNA hybrid RNA and 0.2–0.4 pmol of editosomes. Samples were incubated at 27°C for 3 h after which 15 fmol of two size-standard oligoribonucleotides (FAM_High44 and HEX_Low11) were added. Reactions were stopped by phenol/chloroform extraction and the processed RNAs were EtOH precipitated. Samples were immediately centrifuged for 30 min at 13 000 rpm (4°C) and RNA pellets were washed (70% (v/v) EtOH), dried in vacuo and resolved in 100 μl of Hi-Di™ formamide. Samples were heat denatured at 95°C for 2 min, snap-cooled and separated by capillary electrophoresis (CE) at 12 kV for 25 min using a POP-6 polymer (Thermo Scientific). ‘One-pot’ U-insertion/U-deletion reactions were assembled by annealing of the U-insertion and U-deletion pre-mRNA/gRNA-hybrid RNAs in separate reaction vials. Equimolar amounts (0.2 pmol) of each hybrid RNA were then combined with 0.2–0.4 pmol of editosomes in EB in the presence of 0.5 mM ATP, 0.1 mM UTP and 0.5 mM DTT in a final volume of 30 μl. All incubation and processing steps of the one-pot samples were performed as above. Down-scaled U-insertion and U-deletion assays were performed with 10 fmol of pre-mRNA/gRNA hybrid RNAs and 2.5 fmol of editosomes in a volume of 4 μl. The reduced amounts of material made it possible to skip the phenol extraction and EtOH-precipitation steps of the work flow and enabled a direct sample application onto the CE-system. We also confirmed that the FIDE-assay can be performed in a non-precleaved fashion. For that we used the pre-mRNA mimicking oligoribonucleotide HEX_mRNAdel in conjunction with gRNAdel.RNA-editing reactions were assembled in a 30μl volume in editing buffer /∂noise . The data were baseline corrected and individual peaks were assigned by comparison to the unedited peak of a mock reaction using the electrophoretic migration times of two 11nt- and 44nt-long calibration oligoribonucleotides (FAM_High44 and HEX_Low11) as reference points. Peaks were integrated using a Riemann sum approximation to derive RNA-editing activity values (EA) defined as the ratio of the peak area (A) of the fully edited product (AFE) versus the sum of all peaks, i.e. the unedited (AUE), partially edited (APE) and fully edited (AFE): EA = AFE/∑. EA-values divided by the mass of editosomes represent the specific RNA-editing activity of the editosome preparation in EA/ng. By computing the ratio between the area of each peak and the total integrated area, the fraction of each reaction intermediate was calculated. In the case of 'one-pot' RNA-editing assays, raw data from the electropherograms were first imported into ShapeFinder (Z') as Z' = 1 – (3∂control+ + 3∂control–)/|μcontrol+-μcontrol-|  (Raw data from the CE-runs (relative fluorescence units (RFU) and electrophoretic migration times in seconds) were converted into tab-delimited text files and imported either into OriginPro v8.5 (OriginLab Corporation) or MultiGauge v3.0 . The quality of the CE-separations was assessed by evaluating the signal-to-noise (S/N) ratio as S/N = (μpeFinder  for basecontrol) .A260) using a thermoelectrically controlled UV-spectrophotometer. The temperature was scanned at a heating rate of 1°C/min at temperatures between 20 and 90°C. Absorbance values were recorded with an average time of 0.5 s and data were collected every 0.1°C. Samples contained 1 μM pre-mRNA/gRNA hybrid RNA in 10 mM Na-cacodylate pH 6.8, 65 mM NaCl. Half-maximal melting temperatures (Tm) were calculated from first-derivative plots of absorbance versus temperature δA260/δT = f(T) (Absorbance versus temperature profiles (melting curves) of the different pre-mRNA/gRNA hybrid RNAs were recorded at 260 nm (T = f(T) .The RNase-sensitivity of unmodified and phosphorothioate-modified 5′-Cl oligoribonucleotides  was tested by incubation of 0.4pmol RNA-oligonucleotide with 0.2 pmol of editosomes in 30 μl EB for 3 h at 27°C. Samples were processed and analyzed as described above.in vitro RNA-editing assay (trans-acting gRNAs are mimicked by short oligoribonucleotides (13–22nt) -labeled to enable a radioactive readout of the assay. To convert the system into a fluorescence-based assay we covalently attached different fluorogenic side groups to either the 5′- or 3′-terminal phosphate groups of the two Cl-oligoribonucleotides. 5-Carboxytetramethylrhodamine , 6-carboxy-fluorescin  and 6-hexachlorofluorescein  were chosen as fluorophores. The coupling was performed post-synthetically using a 6-carbon atom spacer to minimize sterical clashes between the fluorophore moieties and the RNA-backbone. Since RNA-modifications can alter the physico-chemical properties of polynucleotides especially in the case of highly hydrophobic side-groups, we analyzed whether the different fluorophores affect the formation of the pre-mRNA/gRNA–hybrid RNAs. Antiparallel base-pairing between the pre-mRNA Cl-fragments and the corresponding gRNAs represents the first step in both in vitro editing reactions and the formation of the bi- and trimolecular RNA-hybrid molecules can be scrutinized by native polyacrylamide gel-electrophoresis. As shown in Figure Tm-values of 68 ± 1.5°C and 47 ± 1.5°C for the U-insertion complex and 64 ± 1.5°C and 55 ± 1.5°C for the U-deletion hybrid. Importantly, all fluorophore-substituted versions of the two trimolecular RNA-complexes show qualitatively and quantitatively identical profiles. The derived Tm-values deviate ≤1°C from the non-fluorophore substituted counterparts and as before, this behaviour is affected neither by the chemical nature nor by the structural positioning of the fluorophore moieties. For a complete list of all measured Tm-values see The established U-insertion/U-deletion ng assay  represenleotides 3–22nt  as well as single turnover conditions (editosomes ≥ RNA-substrate). This enabled us to identify the products of the catalytic conversion as well as all intermediates and side-products. After the reaction, samples were analyzed by automated capillary electrophoresis (CE) coupled to a laser-induced fluorescence (LIF) readout. Figure T. brucei genetic backgrounds. The data were analyzed in a bivariate correlation analysis to calculate Pearson's correlation coefficients (p). Figure p-values ≥0.99 and biological replicates by p-values ≥0.89 demonstrating a high level of experimental robustness. A comparison of the fluorophore CE/LIF-based in vitro assays with the standard radioactive, slab gel-based assays resulted in p-values ≥0.86  a dual fluorophore-labeling approach of the two pre-mRNA-mimicking oligoribonucleotides and (ii) a side-by-side analysis of both, U-insertion and U-deletion editing in one reaction vial. Representative examples of the generated data-sets are shown in Figure in vitro reactions can be downscaled to tailor the assay for high-throughput screening applications. This was done in a stepwise fashion using the signal to noise ratio (S/N) of the CE-electropherograms as a proxi. As lower limit we determined 2.5–10 fmol of each, editosomes and fluorophore-modified pre-mRNA/gRNA-hybrid in a reaction volume of 4μL. At these conditions signal intensities remained ∼300-fold over background for high intensity peaks and 50-fold over background for low intensity peaks. Because of the minute amounts of protein and RNA in the samples, reactions were stopped by directly adding formamide thereby eliminating the time-consuming phenol and EtOH-treatment steps. Furthermore, the small assay volume enables the usage of multiwell plates, which makes the test suitable for HTS-experiments. As a statistical HTS-quality score  the ribonucleolytic stability of the substrate RNAs and (ii) possible sequence context effects because the U-deletion and U-insertion substrate RNAs are of different sequence. Since editosomes are purified from whole cell lysates co-purifying and/or contaminating RNases can be challenging e.g. see . The prod Figure . Phosphod Figure . Incubatd Figure . While ud Figure , howeverd Figure . Finally3-UTP, 5-Br-UTP, 5-OH-UTP, 4-S-UTP, 2-S-UTP, 5-biotin-UTP, 5-CH3N3-UTP etc.) as well as ribose-modified analogs such as 2′-CH3O-UTP, 2′-NH2-UTP, 2′-F-UTP, 2′-N3-UTP and Ara-UTP were tested. A list of all compounds is shown in Figure 2+- and Mn2+-cations in the reaction. Altogether 26 samples were tested, which on a 48 multicapillary CE/LIF-instrument can be assayed in just 1.5h generating 260 quantitative data points on the formation of the fully edited product and on every reaction intermediate. As shown in Figure 3-UTP (+4Us) and 2-S-UTP (+4 and +5U′s). Mn2+-ions have an enhancing effect, which can result in the incorporation of otherwise non-inserted UTP-analogs. Together, the data demonstrate the versatility of the FIDE-in vitro system by enabling a comprehensive and quantitative inhibitor screening in a very short time.To demonstrate the high-throughput potential of the FIDE-assay, we performed a small-scale, proof-of-principle screening experiment. For that we made use of the non-natural U-insertion pre-mRNA/gRNA-substrate described above and examined whether U-nucleotide analogs might interfere with the U-insertion reaction. Both, pyrimidine ring-modified compounds (5-CHin vitro assay to monitor editosome function in African trypanosomes and related organisms into a high-throughput analysis format. Core attribute of the revised assay is the usage of fluorophore-labeled RNA-editing substrate RNAs. This enables the automated electrophoretic separation of the products of the catalytic conversion using state-of-the-art, high-throughput capillary electrophoresis instruments coupled with laser-induced fluorescence readout systems. As such the assay generates quantitative data of all educts, intermediates and products and by using multicapillary instruments it can be performed in a highly parallel format. We optimized the assay by downscaling the required material quantities as well as the reaction volume to adapt the procedure to all available multiwell-plate formats. Additional improvements include the usage of non-natural, RNase-resistant RNA-substrates, which enable large-scale screening experiments using crude mitochondrial extracts instead of highly purified editosome preparations. We also verified that the experimental setup tolerates multiplex-type, fluorophore-labeling strategies, which expands the parallel analysis-capacity of the assay further. As demonstrated in a pilot screening experiment, the assay is applicable to larger sample-size studies to probe specific mechanistic aspects of the RNA-editing reaction. However, the assay is especially well suited to conduct large-scale, high-throughput screening experiments to identify small molecule inhibitors. Since RNA-editing is an essential biochemical pathway in African trypanosomes, an assay format that permits a robust, quantitative as well as time- and material-resourceful analysis should be helpful in the search for novel therapeutics to challenge African trypanosomiasis and related diseases.In summary, we have converted the standard gkaa658_Supplemental_FileClick here for additional data file.


Betanecrovirus, family Tombusviridae) contains several well studied regulatory RNA elements. Here, we explore a previously unexamined region of the viral 3′UTR, the sequence located upstream of the 3′-cap independent translation enhancer (3′CITE). Our results indicate that (i) a long-range RNA–RNA interaction between an internal RNA element and the 3′UTR facilitates translational readthrough, and may also promote viral RNA synthesis; (ii) a conserved RNA hairpin, SLX, is required for efficient genome accumulation; and (iii) an adenine-rich region upstream of the 3′CITE is dispensable, but can modulate genome accumulation. These findings identified novel regulatory RNA elements in the 3′UTR of the TNV-D genome that are important for virus survival.RNA elements in the untranslated regions of plus-strand RNA viruses can control a variety of viral processes including translation, replication, packaging, and subgenomic mRNA production. The 3′ untranslated region (3′UTR) of Tobacco necrosis virus strain D (TNV-D; genus  Tobacco necrosis virus strain D (TNV-D), and other plus-strand RNA plant viruses contain RNA elements throughout their genomes that regulate different viral processes ,5,6,7,8.Tombusviridae, and is the type-member of the genus Betanecrovirus, which also includes Beet black scorch virus (BBSV) . Ho. Ho1]. HTombusvirus genomes do not contain any functionally important secondary structures in the region immediately upstream from their DL sequences. In contrast, TNV-D is predicted to contain two conserved hairpins in this location. SLY is the first predicted RNA secondary structure downstream of the p29 stop codon in the TNV-D genome C. ModifiSLX is also conserved in both betanecroviruses C. ModifiSome tombusvirids, in addition to their 3′CITE, require an interaction with PABP to enhance translation . In the This study provides an initial assessment of the functional relevance of RNA elements in the upstream third of TNV-D’s 3′UTR. The results revealed the presence of RNA elements that were either similar to other tombusvirids (UL–DL interaction) or unique to the betanecrovirus genus (SLX and SLY). Among the four RNA elements investigated, SLX and the DL  were most functionally relevant. The UL–DL interaction promotes readthrough, and may facilitate RNA replicase complex assembly , whi


Small RNA sequencing showed the two base pair deletion in mir390 substantially interfered with miR390 biogenesis. In Arabidopsis, CRISPR/Cas9‐mediated editing of the miR160* strand confirmed that the asymmetric structure of miRNA is not a necessary determinant for secondary siRNA production. CRISPR/Cas9 with double‐guide RNAs successfully generated mir160a null mutants with fragment deletions, at a higher efficiency than a single‐guide RNA. The difference between the phenotypic severity of miR160a mutants in Col‐0 versus Ler backgrounds highlights a diverged role for miR160a in different ecotypes. Overall, we demonstrated that TALENs and CRISPR/Cas9 are both effective in modifying miRNA precursor structure, disrupting miRNA processing and generating miRNA null mutant plants.MicroRNAs (miRNAs) are 20‐24 nucleotides (nt) small RNAs functioning in eukaryotes. The length and sequence of miRNAs are not only related to the biogenesis of miRNAs but are also important for downstream physiological processes like ta‐siRNA production. To investigate these roles, it is informative to create small mutations within mature miRNA sequences. We used both TALENs (transcription activator‐like effector nucleases) and clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR‐associated protein 9 (Cas9) to introduce heritable base pair mutations in mature miRNA sequences. For rice, TALEN constructs were built targeting five different mature miRNA sequences and yielding heritable mutations. Among the resulting mutants,  The biogenesis of miRNAs was thoroughly characterized in recent years (reviewed in Achkar MIRNA (MIR) gene by RNA polymerase II (Pol II), yielding a long, polyadenylated, single‐stranded RNA. This RNA forms a hairpin structure and is processed by DCL1 (DICER‐LIKE 1), a member of the RNase III family. The product of this process is a much shorter RNA with a stem‐loop structure named the pre‐miRNA. The pre‐miRNA stem‐loop structure usually exposes a duplex of the mature miRNA and miRNA* on the stem, allowing DCL1 to cleave the duplex out of the stem loop  are another pathway unique to plants. A ta‐siRNA precursor gene is transcribed into long, polyadenylated mRNA and subsequently processed into ‘phased’ siRNAs (characterized by precise spacing of 21 or 24 nt); the ta‐siRNAs inhibit gene expression by binding to complementary mRNA sequences. The phased processing and spacing of ta‐siRNAs derive from initiation by a miRNA ‘trigger’ and generation of a double‐stranded precursor by RDR6 (RNA‐DEPENDENT RNA POLYMERASE 6). It was reported that a 22‐nt length of the miRNA trigger is crucial to produce ta‐siRNA  and CRISPR/Cas ‐based genome editing technologies are promising tools to precisely alter any genomic loci of interest in eukaryotes  was dominantly produced, while the wild‐type MIR398b produced mostly a 21‐nt miRNA  as wild type: heterozygous: ungerminated seeds. Random genotyping of ungerminated seeds indicated that they were all homozygous mutants of mir390. The homozygous mutant seeds could either produce a root without any shoot (i.e. a SAM defect) or not germinate at all due to lack of embryonic shoot apical meristem (SAM)  callus line. mir390−/− calli were screened and selected using the two‐step PCR method described above. The rice MIR390 gene was PCR‐amplified as a genomic fragment and cloned into the pCambia1300 vector for complementation of mir390−/− mutant through rice transformation. The pCambia1300/MIR390 plasmid was then transferred into mir390−/− callus cells. As previously mentioned, mir390−/− showed a shootless phenotype. However, the transformed calli could be regenerated into plants with normal shoots, a phenotype that was rescued by the genomic fragment of MIR390. The sequencing results confirmed the shooting calli were indeed mir390−/− mutants containing a MIR390 transgene  rice mutant, which is mutant in the orthologous gene to AGO7 in Arabidopsis. Although AGO7 and miR390 are demonstrated in Arabidopsis to be highly selective to each other, the different phenotypes of shl4 and ago7 in these two plant species raise questions of how they interact with miR390 differently in rice and Arabidopsis , which would generate an asymmetric structure on the miR160a precursor and resultant miR160a/miR160a* duplex   showed a mixed Sanger sequencing result Figure a, sugges) Figure a. The li1 Figure b. Consis1 Figure c. We alss Figure d. Moreovmir160a*‐Δ5 is a null mutant that is caused by the secondary structure change, we identified an additional PAM site on the miR160a strand, 49‐bp upstream of the aforementioned miR160a* PAM site  are serrated . CRISPR/Cas9 can be used to create miRNA mutants either on the miRNAs with appropriate PAM sequences, or delete the whole region by targeting both ends of the stem‐loop sequence, but internal mutations may not be possible. Deleting the whole stem loop of a miRNA should yield a null mutant, which may be useful for some analysis, such as assessing miRNA function or targets  using the criteria for TALEN target selection as previously described  35S promoter. The TALEN‐R was similarly assembled and ligated into the same pCambia1300 plasmid for expression under promoter of the maize UBIQUITIN 1 gene. The pCambia1300 plasmid bearing both TALEN‐L and TALEN‐R was then transferred into the rice variety Kitaake (Oryza sativa subsp. japonica) to regenerate stable transgenic plants through Agrobacterium‐mediated transformation.TALEN constructs were assembled as previously described  Cas9 gene under the Arabidopsis UBQ10 gene promoter, as previously described  or mutant mir390 (MIR390‐F3). Combined with a common reverse primer in the first step, both pairs of primers were used to amplify the corresponding alleles of MIR390. The first step PCR increases template concentration and allows more specific PCR at the second step. By the end, MIR390‐F2 gave only positive signal when there was wild‐type allele and MIR390‐F3 only functioned in the presence of a mutant allele. Genotypes  were determined by analysing the amplicons of the second step PCR.A two‐step PCR method was used to genotype MIR390 was PCR‐amplified with primers miR390‐F1 and miR390‐R1 (Table MIR390 genomic fragment into a modified pCambia1300 vector at the restriction cloning sites (EcoRI and HindIII) for transformation of mir390−/− derived callus cells.Kitaake genomic DNA was used as PCR template for high‐fidelity PCR. The genomic region of R1 Table . The PCRΔΔCt method . One μg of total RNA was then treated with DNase I at 37 °C for 20 min, followed by heat inactivation with EDTA. The DNA‐free RNA was then reverse‐transcribed using SuperScript III (Thermo Fisher Scientific), followed by quantitative PCR using primers as listed in Table et al., et al., GSE130510 for Arabidopsis and GSE130239 for rice.Total RNAs were prepared from plant tissues using a TRIzol reagent‐based method. The plant materials were homogenized and reconstituted in TRIzol solution. The solution in turn was extracted by chloroform. Isopropanol was used to precipitate the total RNA. DEPC‐treated water was used to dissolve RNA. The total RNA underwent miRNA isolation using Qiagen miRNA purification kit according to the manufacturer’s instruction. Small RNA libraries were made using 5 µg total RNA with NEBNext Small RNA Library Prep Set as described (Mathioni The authors declare no conflict of interest.H.B., B.Y., Q.F. and B.C.M. conceived and designed the experiments. H.B., Q.F., R.L. and B.L. performed the research. R.X. and S.N.C. analysed the data. H.B. and Q.F. wrote the manuscript. B.C.M. and B.Y. revised the manuscript with input from other authors.Figure S1 Analysis of the mir160a mutants by CRIPSR/Cas9 with single‐guide RNA.Figure S2 Characterization of the mir160a mutant with 5‐bp deletion.Figure S3 Agarose gel analysis of the PCR products from CRISPR/Cas9‐transformed plants with double‐guide RNAs.Table S1 Primers and synthesized DNA sequences used in this study.Click here for additional data file.Table S2 Small RNA library data for miRNA mutants.Click here for additional data file.


Next, we employed small-angle X-ray scattering techniques to study solution conformation and low-resolution structures of these RNAs, which suggest that the 3’ terminal regions are highly extended as compared to the 5’ terminal regions for both viruses. Using computational modeling tools, we reconstructed 3-dimensional structures of each RNA fragment and compared them with derived small-angle X-ray scattering low-resolution structures. This approach allowed us to reinforce that the 5’ terminal regions adopt more dynamic structures compared to the mainly double-stranded structures of the 3’ terminal regions.Viral infections are responsible for numerous deaths worldwide. Flaviviruses, which contain RNA as their genetic material, are one of the most pathogenic families of viruses. There is an increasing amount of evidence suggesting that their 5’ and 3’ non-coding terminal regions are critical for their survival. Information on their structural features is essential to gain detailed insights into their functions and interactions with host proteins. In this study, the 5’ and 3’ terminal regions of Murray Valley encephalitis virus and Powassan virus were examined using biophysical and computational modeling methods. First, we used size exclusion chromatography and analytical ultracentrifuge methods to investigate the purity of  Family Flaviviridae consists of small, positive-sense single-stranded RNA viruses that replicate within host cells of arthropods and/or vertebrates. Flaviviruses include deadly viruses, such as Murray Valley encephalitis virus (MVEV), Powassan virus (PowV), Japanese encephalitis virus, dengue virus, Zika virus, West Nile virus and yellow fever virus. The World Health Organization and the Centers for Disease Control both cite flaviviruses as a global health threat owing to the ease of transmission by mosquitoes, and the lack of efficient therapeutic or immunoprophylactic strategies . There iCulex annulirostris) and birds [MVEV is a member of the Japanese encephalitis serological complex of flaviviruses and was first isolated in 1951 during the initial outbreak in Australia [nd birds ,7. The lnd birds . There hnd birds . PowV isnd birds . There hnd birds . It was nd birds  and has nd birds . The incFlaviviral genomes consist of a single-stranded RNA molecule approximately 10-11k nucleotides long, depending on species. The RNA genome contains one single open reading frame (ORF), which is flanked by terminal regions (TRs) . The flaAlthough several attempts have been made to study RNA structures in general ,34, detaWe prepared cDNA sequences under the control of a T7 RNA polymerase promoter with two additional G nucleotides on the 5’ end followed by an XbaI restriction enzyme cut site (T^CTAGA). We designed our constructs for MVEV and PowV based on the Genebank sequences of KX229766.1and EU670438.1, respectively. All RNA constructs used in the experiments are listed as follows:MVEV 5TR 1-96nt5’GGAGACGUUCAUCUGCGUGAGCUUCCGAUCUCAGUAUUGUUUGGAAGGAUCAUUGAUUAACGCGGUUUGAACAGUUUUUUGGAGCUUUUGAUUUCAAU3’MVEV 3TR 10914-110145’GGCCUGGGAAAAGACUAGGAGAUCUUCUGCUCUAUUCCAACAUCAGUCACAAGGCACCGAGCGCCGAACACUGUGACUGAUGGGGGAGAAGACCACAGGAUCUU3’POWV 5TR 1-1115’GGAGAUUUUCUUGCACGUGUGUGCGGGUGCUUUAGUCAGUGUCCGCAGCGUUCUGUUGAACGUGAGUGUGUUGAGAAAAAGACAGCUUAGGAGAACAAGAGCUGGGAGUGGUUU3’POWV 3TR 10735-108395’GGCCCCCAGGAAACUGGGGGGGCGGUUCUUGUUCUCCCUGAGCCACCACCAUCCAGGCACAGAUAGCCUGACAAGGAGAUGGUGUGUGACUCGGAAAAACACCCGCUU3’2). Next, we analyzed the pooled fractions using Urea-Polyacrylamide Gel Electrophoresis (Urea-PAGE). We mixed 10 µL of ~500 nM RNA with 2 µL of RNA-loading dye and placed them into a 1.0 cm well PAGE casting plate (Bio-Rad Laboratories (Canada), Mississauga, ON). Finally, we ran the Urea-PAGE (7.5%) at 300V at room temperature for 25 minutes in 0.5 X TBE, followed by staining with Sybr Safe  and UV visualization. Prior to performing analytical ultracentrifugation (AUC) and small-angle X-ray scattering (SAXS) experiments, we heated RNA to 95 °C for 5 min and cooled passively to room temperature to facilitate refolding.We performed in vitro transcription reactions using T7 RNA polymerase to prepare each RNA, followed by their purification using a Superdex 200 increase  on an ÄKTA pure FPLC (fast protein liquid chromatography) system  at 0.5mL/min. We collected and pooled fractions containing purified RNAs from the size exclusion chromatography (SEC) fraction collector. Pooled fractions were ethanol precipitated and resuspended in RNA buffer  - AUC data for FPLC-purified RNA samples were collected using a Beckman Optima AUC centrifuge and an AN50-Ti rotor at 20 °C. We loaded MVEV 5’ (0.6 uM), MVEV 3’ (0.5 uM), PowV 5’ (0.6 uM) and PowV 3’ (0.58 uM) samples into Epon-2 channel centerpieces in 50 mM Tris pH 7.5, 100 mM NaCl, and 5 mM MgCl package , followe package  and gene package . We deri package  and esti−1 was injected into a buffer equilibrated Shodex KW403-4F  size exclusion column at a flow rate of 0.160 mL per minute. Each frame was exposed to the X-rays for 3 seconds. Each sample peak region was buffer subtracted and merged using Primus [gR) and study homogeneity of samples [P(r)) analysis using the program GNOM [gR and the maximum particle dimension  dependence approaches zero). We used information from the P(r) plot to generate the models using DAMMIN [We utilized the B21 beamline at Diamond Light Source  to collect small-angle X-ray scattering (HPLC-SAXS) data as previously described . Making g Primus  or ScÅttg Primus  as descrg Primus . We anal samples . We also samples  to inves samples . Next, wram GNOM , which pg DAMMIN , with nog DAMMIN . Lastly,g DAMMIN , as descg DAMMIN ,51.gR and the goodness-of-fit parameter (2χ) [2χ values and aligned them with low-resolution structures using the program SUPCOMB [To generate secondary structure predictions used downstream, we used MC-Fold  to prediter (χ2) . Finally SUPCOMB . We purified the in-vitro transcribed RNAs using a Superdex 200 increase column connected to the ÄKTA FPLC unit. −13 seconds, s values corrected to 20 °C for water), as summarized in w of ~38 kDa have a slightly higher sedimentation coefficient compared to the MVEV TRs with Mw of ~32 kDa ) vs. (2q)), which aids detection of purity and allows determination of the gR (average root mean squared radius from the center of the mass for a biomolecule) from the data belonging to the low-q region [q data demonstrates that all four RNAs are monodispersed and devoid of any aggregation. Based on the Guinier analysis, we obtained gR values of 29.01 ± 0.22 Å, 45.72 ± 0.48 Å, 34.65 ± 0.19 Å and 35.84 ± 0.13 Å for MVEV 5’ TR, MVEV 3’ TR, PowV 5’ TR and PowV 3’ TR respectively (see I(q)/I(0)*(q*gR)2 vs q*gR) that allow detection of the folding state of biomolecules [q*gR = 1.73 [Small-angle X-ray scattering allows for structural determination of biomolecules and their complexes under physiological conditions, albeit at low-resolution. Recent improvements in instrumentation employ an HPLC unit connected in line with SAXS detection to improve the monodispersity of samples and to resolve species of interest from aggregated and degraded products ,58. Thisq region . Figure olecules ,58. For ln(I(q)) vs. (q), P(r), gR and maxD (maximum particle dimension) for all four RNAs. It is important to note that unlike Guinier analysis, which is restricted to the data in the low-q region, the P(r) analysis utilizes a wider-range of the dataset and aids in a reliable determination of gR and maxD. As outlined in P(r) analysis, we obtained the maxD of 100 Å, 150 Å, 105 Å, and 130 Å for MVEV 5’ TR, MVEV 3’ TR, PowV 5’ TR, and PowV 3’ TR respectively. Furthermore, we obtained the gR values of 30.16 ± 0.11 Å, 46.16 ± 0.18 Å, 34.48 ± 0.06 Å and 38.39 ± 0.08 Å for MVEV 5’ TR, MVEV 3’ TR, PowV 5’ TR and PowV 3’ TR respectively. These values are very similar to those obtained from Guinier analysis, which indicates that the data are suitable for low-resolution shape reconstruction. The shape of the P(r) plot is indicative of the solution conformation of biomolecules. For example, we would have expected a bell-shaped P(r) distribution curve with a maximum at ~maxD/2 [Next, using program GNOM , we perf ~Dmax/2  for a glP(r) data . We calculated a total of 12 models for 5’ and 3’ TRs of MVEV and PowV and noted that individual models had an excellent agreement between the experimentally obtained scattering data and the calculated scattering data. The χ2 values in each case are ~0.8, which represents an agreement between the experimentally collected scattering data and the low-resolution model derived scattering data  2 value. We also determined gR from MC-Sym derived structures. gR  and χ2 values  for each RNA system, which suggests that the RNA molecules under investigation can adopt a variety of conformations. For example, in the case of 5’ TR of MVEV, the distribution of gR values range from 30 ± 5 Å, whereas the experimentally determined gR for this construct is 30.16 Å. Similarly, for PowV 3’ TR, experimentally determined gR is 38.39 Å, which largely aligns with the distribution of gR values of MC-Sym derived structures. On the other hand, the MC-Sym derived models for MVEV 3’ TR mainly under-represent the gR values when compared to experimental gR of 46.16 Å. In contrast, for PowV 5’TR, Mc-Sym derived structures have a higher gR distribution (~45Å) compared to the experimentally determined gR (34.48 Å). Moreover, as presented in 2 values for MC-Sym derived 3’ TR of MVEV and PowV structures have better distribution around 1.5 χ2, compared to MVEV and PowV 5’ TRs. SAXS is an outstanding method that allows the determination of solution structures of biomolecules, albeit at low-resolution. Techniques such as X-ray crystallography and nuclear magnetic resonance spectroscopy provide high-resolution structural information but have limited applications for RNA structural studies due to challenges obtaining high-quality crystals and RNA-labeling, respectively. An alternative to this approach is to employ computational modeling of RNA structures and screen those structures using experimental data. We employed the MC-Fold/MC-Sym pipeline developed by Parisien and Major . For eac2 values to further investigate if they align with the low-resolution averaged filtered models for each RNA system. 2 fit MC-Sym derived structures for MVEV and PowV, indicating that these structures agree well with the low-resolution models based on SAXS data. Overall, MC-Sym derived structures for the 3’ TRs of MVEV and PowV . Furthermore, the S values for 5’ and 3’ TRs of PowV are 4.50 S and 4.53 S. This similarity correlates with the similarities of their wM of ~ 37 kDa , which are highly similar to those calculated by means of P(r) analysis (P(r) distribution reveals a quick increase to the highest point and then a gradual decrease where the electron-pair-distance approaches zero. A globular molecule would result in a Gaussian-like distribution, which is not evident in any of the RNA, and suggests their elongated nature. The most extended molecule is MVEV 3’ TR resulting in a maxD of 150 Å, even though its molecular weight is 31.57kDa . A similar trend was also observed for gR for both 5’ TRs where the MC-Sym derived structures displayed a wider distribution of Rg, suggesting that these RNAs could adopt multiple conformations. The correlation between gR and 2χ was far more pronounced in the more elongated TR RNAs - PowV 3’ TR and MVEV 3’ TR . Howeve


Deoxyribozymes (DNAzymes) with RNA hydrolysis activity have a tremendous potential as gene suppression agents for therapeutic applications. The most extensively studied representative is the 10-23 DNAzyme consisting of a catalytic loop and two substrate binding arms that can be designed to bind and cleave the RNA sequence of interest. The RNA substrate is cleaved between central purine and pyrimidine nucleotides. The activity of this DNAzyme in vitro is considerably higher than in vivo, which was suggested to be related to its divalent cation dependency. Understanding the mechanism of DNAzyme catalysis is hindered by the absence of structural information. Numerous biological studies, however, provide comprehensive insights into the role of particular deoxynucleotides and functional groups in DNAzymes. Here we provide an overview of the thermodynamic properties, the impact of nucleobase modifications within the catalytic loop, and the role of different metal ions in catalysis. We point out features that will be helpful in developing novel strategies for structure determination and to understand the mechanism of the 10-23 DNAzyme. Consideration of these features will enable to develop improved strategies for structure determination and to understand the mechanism of the 10-23 DNAzyme. These insights provide the basis for improving activity in cells and pave the way for developing DNAzyme applications. Deoxyribozymes or DNAzymes are single-stranded DNA molecules that are capable of catalyzing a variety of chemical reactions, including RNA cleavage  and ligaIn this review, we aim at providing an overview of the structural and functional data on the RNA-cleaving 10-23 DNAzyme. First, we highlight the potential of the 10-23 DNAzyme for therapeutic applications. We then summarize efforts to gain structural insights on DNAzymes utilizing various methods and point out the challenges for this endeavor. We further describe the reaction mechanism with its relevant parameters. In the following sections, we take a closer look at the functional data available on the impact of nucleotide substitutions within the binding arms as well as substitutions and modifications within the catalytic loop. Finally, we outline the influence of various metal ions on activity.The ability of the 10-23 DNAzyme to cleave disease-related (messenger or non-coding) RNA makes it particularly attractive as a therapeutic agent. It is highly versatile yet specific. In comparison to the currently more established approaches for post-transcriptional gene silencing, such as the use of RNA interference (RNAi) or antisense deoxyribonucleotides (AS-ODNs), DNAzymes have several remarkable advantages. These advantages include the fact that they—supposedly—are self-sufficient biocatalysts that does not rely on the presence of other biomolecules the way that RNAi relies on argonaut proteins and AS-ODNs on the action of RNase H. These features make DNAzymes also perfect candidates to target RNA-based viruses such as human immunodeficiency viruses (HIV) ,28,29,30Structural information on DNAzymes is scarce and the absence of detailed information on the spatial arrangement and the metal coordination sites prevent a deeper understanding of the molecular mechanism of DNAzyme catalysis. Based on its similarity to the hammerhead ribozyme, the RNA-cleaving reaction is believed to involve a transesterification  32,33].,33.32,33All attempts described so far have focused on determining the structure using X-ray crystallography. However, the formation of highly ordered nucleic acid crystals is often challenging due to the surface properties of nucleic acids. While the surfaces of proteins consist of hydrophobic patches that favour intermolecular interactions, the surfaces of nucleic acids are dominated by negatively charged and regularly ordered phosphate groups. Uniform surfaces prevent the formation of distinct interactions between molecules in the crystal lattice and lead to the formation of crystals with poor long-range order. In addition to the challenge of obtaining high quality crystals, solving the phase problem is an additional obstacle in nucleic acid crystallography. The phase problem arises from the loss of phase information during the diffraction experiment. When collecting X-ray diffraction data from a crystal, the intensities of the diffracted waves scattered from a series of planes are measured . From thThe catalytic activity of the 10-23 DNAzyme is shown in the scheme of ing with B. The thThe combined reactions –3) can  can 3) cligation . Under tferences ,53,54,55t, the rate constantUnder single-turnover conditions, that is, DNAzyme is in excess over RNA or the reaction is started by addition of The 10-23 DNAzyme cleaves RNA substrates with high nucleotide selectivity at purine-pyrimidine junction sites (rRrY) in the presence of divalent metal ions. The purine nucleotide of the RNA cleavage site remains unpaired while the pyrimidine nucleotide forms a base pair with one of the deoxyribonucleotides of the DNAzyme. The pyrimidine nucleotide of the RNA cleavage site as well as the paired base of the DNAzyme will be referred to as being in position +1 in the following because they are located “downstream” of the actual site of cleavage, that is, between the purine and pyrimidine nucleotide see . In this+1) for efficient cleavage rates. This is provided by standard nucleotides when followed by uridine  substitutions within the binding arms of the 10-23 DNAzyme have shown that modifications that promote an A-form helix of the binding arms enhance the activity of the 10-23 DNAzyme [Furthermore, studies with 2′- DNAzyme ,63. Whil DNAzyme .Taken together, a certain degree of flexibility in the base pairing between the substrate nucleotides next to the cleavage site and the corresponding deoxynucleotides in the DNAzyme binding arms is crucial for cleavage activity of the 10-23 DNAzyme. The required conformational freedom might be a hint for a significant rearrangement of the RNA cleavage site towards functional deoxynucleotides in the catalytic loop sequence. This assumption is further supported by the observation that the transition from the catalytic loop to the substrate binding arm, that is, the connection between dAThe structure of the DNAzyme in complex with its RNA substrate in a catalytically relevant conformation remains unknown. Therefore, much effort has been spend into determining how the deoxynucleotides within the catalytic loop affect DNAzyme function. Several studies have investigated the role of each individual nucleobase ,68,69,702. They used the total yield of the reaction product after 20 min as an indicator of DNAzyme activity. This approach is less labor-intense, however, the results are less reliable as compared to kinetic measurements. Reference [kobs values compared to the unmodified DNAzyme in the following order: dakobs values significantly further compared to the substitution by an abasic deoxynucleotide. In case of daYReference  analyzedeference  found theference  found theference  were perThe dramatically reduced DNAzyme activity after replacement of dGinactive , emphasifunction   in the hydrolysis reaction. With regard to the importance of the 6-keto group in the proximal deoxynucleotide dGP as well as the 6-keto group of dGP-chirality also suggested that the RP is directly involved in metal ion coordination. Since significant thio and rescue effects were also detected for the oxygen atoms at positions d2P, d4P, d10P, d11P, d12P, and d13P, these ligands may also be involved in the coordination of metal ions [At position dAactivity . Replaceactivity . In contactivity . These factivity . Studiesactivity  revealedactivity  analyzedeoisomer . The acteference  performeeference  proposestal ions .R)- and (S)-2′-C-methyl-purine as well as locked conformations had a negative effect on the catalytic activity of the DNAzyme [Thymidine dTuding dI , an abasuding dI  or a comuding dI . It has ancement . Introduactivity . The wea DNAzyme ,80. Howeubstrate . Activitactivity .YYMutagenesis analyses by Reference  with subomal RNA . Albeit atalysis ,70,72.To analyze the effect of DNAzyme sequence on its RNA cleavage activity, Reference  removed  or Mn2+ . Since a ligands .A summary of the effects of substitutions and modifications of single deoxynucleotides and functional groups on the activity of the 10-23 DNAzyme is given in Ka value or the nucleophilic character of a water molecule. They are also able to alter the properties of functional groups by polarization. Ka values of coordinating water molecules, and the affinity to functional groups tremendously affect the catalytic properties of the metal ion. Studies on the effect of metal ions on the DNAzyme activity have shown that divalent metal ions promote the reaction in the following order: The role of metal ions in ribozyme-mediated catalysis ,87,88,89lsewhere ). Metal g2+ ions . Its act2+, Co2+ . These f2+, Co2+ , where ther Mg2+ , Mn2+ [92+ [Mn2+ , or Cd2+2+ [Mg2+  or Mn2+ 2+ [Mn2+ . For allt pH 7.5 .Ka value of different metal ions and they found that the logarithm of the kobs value was inversely proportional to the pKa value. Folding of the DNAzyme in the presence of different Reference  compared domain) . The autibozymes ,99, in DNorth-type ) and 2′-endo (here termed South-type) conformation, the DNA residues of the binding arms and the catalytic loop have the sugar moieties in the 2′-endo conformation, as in B-type DNA. In the presence of South-endo conformation, in the presence of North-endo conformation predominates. Comparing the structures of the native and mutated DNAzymes derived from MD simulations, Reference [Reference  investig helices . While teference  found theference , which seference  reportedeference . Refereneference  has showstrategy . A combiis of α-  and γ-caatalysis , since ted limit ,33. Howeed limit , it is latalysis  as an adatalysis .R)-CHS)-CHR)-CHRP is directly involved in metal ion coordination [SRRThe exhaustive characterization of different 10-23 DNAzyme mutants and chemical modifications provide insights into the impact of various deoxynucleotides and functional groups on DNAzyme catalysis, even in the absence of high-resolution structural information. The mutagenesis studies by References ,70 clear DNAzyme . The mostroduced . The acttroduced . Positiodination . Based odination , a Mg2+ stigated ,103. Herferences ,96. Summferences ,33,103, eference  and Refetalysis) .Yall-RNA substrates other experiments were carried out using chimeric DNA/RNA substrates, with only the nucleotide at the cleavage site being a ribonucleotide. Also, DNAzyme and RNA substrate design can have a massive influence on the kinetic behavior of the DNAzyme, since the length and sequence dramatically influence the dissociation constants of the DNAzyme:RNA complex. In addition, temperature, monovalent cation levels as well as the type of reaction buffer can have an impact on DNAzyme activity. In Although the influence of deoxynucleotide substitutions and modifications, deoxyribose and phosphate modifications within the catalytic loop and the binding arms of the 10-23 DNAzyme, as well as its tolerance towards different metal ions and pH conditions have been extensively studied, the mechanism by which the DNAzyme cleaves its specific RNA substrate as well as the role of different functional groups is only poorly understood. The main cause is arguably the absence of satisfactory structural data of the 10-23 DNAzyme. However, one major obstacle when it comes to the interpretation of different mutational studies is the low level of comparability of the experimental setup. While some experiments were designed to determine only the yield after a given period of time (Efforts to elucidate the reaction mechanism are essential for attempts to use DNAzymes for therapeutic applications. Structural information on DNAzymes is scarce, therefore, we rely on functional data for insights into DNAzyme catalysis. The data summarized in this review provides interesting insights into the role of the different regions of the 10-23 DNAzyme. The binding arms of the DNAzyme are responsible for substrate recognition and form a double helical structure, while the catalytic loop forms an unusual three-dimensional conformation. Taking a closer look at the cleavage site, we conclude that the neighboring nucleotides of the scissile A or G in the RNA and DNA strand play an important role in catalysis. All mutations and modifications at these positions indicate a flexibility or equilibrium of rNStudies focusing on the residues within the catalytic loop have shown that the exocyclic groups of the nucleotides dGThe challenge remains to understand the mechanism of 10-23 DNAzyme catalysis and the role of different functional groups in the DNAzyme sequence. We believe that three aspects are crucial for future studies. First, studies using comparable experimental setups and conditions would be beneficial for the research field. This includes determining rate constants instead of yields and using RNA substrates instead of chimeric substrates. In addition, the effects of buffers and ionic strength should be investigated thoroughly, to ensure that effects observed at different conditions are not a result of the reaction conditions. Second, starting conditions of DNAzyme reactions should be chosen in respect of the desired output. For example, a condition with preformed, buffer-equilibrated 1:1 DNA:RNA complexes and reaction initiation via addition of divalent metal ions should be used to investigate effects of the divalent ions on single-turnover reaction rates, while excess of substrate is essential to characterize the Michaelis-Menten parameters of DNAzyme catalysis. Finally, a high-resolution structure of the 10-23 DNAzyme is necessary to obtain detailed insights into the spatial arrangement of the DNAzyme in complex with its RNA substrate. A major challenge is the formation of irrelevant conformations due to the palindromic sequence. This challenge can be overcome by using variants of the DNAzyme with single nucleotides exchanged in the catalytic loop. Some of these variants retain a considerable level of activity, but the mutation leads to a sequence that is not palindromic and, thus, should not form the biological irrelevant duplex conformation. Another attempt to stabilize the catalytically active form of the complex is to use DNA- or RNA-binding proteins that prohibit duplex formation. Alternatively, methods not relying on crystallization, including NMR and EPR spectroscopy could provide valuable insights into molecular design and dynamics of 10-23 DNAzymes when in complex with a target RNA fragment. Initial experimentally validated structural models will also be essential as reliable starting structure for in silico techniques that may provide more detailed mechanistic insights into the different states of the catalysis process.Taken together, we have summarized molecular features of the 10-23 DNAzyme in this review in order to contribute to a better understanding of these fascinating biocatalysts. Our detailed analysis reveals important aspects to be considered in future attempts to elucidate the structure of DNAzymes in complex with the RNA substrate and unravel the mechanism of the reaction catalyzed by the 10-23 DNAzyme.


Understanding the molecular determinants of retroviral genome packaging is important for drug discovery and development of vectors for gene delivery. We show that the HIV-1 leader, which contains the RNA elements necessary for genome packaging, binds approximately two dozen copies of the cognate NC protein with affinities ranging from ∼40 nM to 1.4 µM. Binding to the four highest-affinity “initial” binding sites occurs with endothermic energetics attributed to NC-induced localized RNA melting. Mutations that stabilize these sites inhibit NC binding in vitro and RNA packaging in transfected cells. A small-molecule inhibitor of RNA packaging binds specifically to the initial NC binding sites and stabilizes the RNA structure. Our findings identify a potential RNA Achilles’ heel for HIV therapeutic development. Kd ≲ 400 nM) reside within a ∼150-nt region of the leader sufficient to promote RNA packaging . The four initial binding sites with highest affinity reside near two symmetrically equivalent three-way junction structures. Unlike the other high-affinity sites, which bind NC with exothermic energetics, binding to these sites occurs endothermically due to concomitant unwinding of a weakly base-paired [UUUU]:[GGAG] helical element. Mutations that stabilize base pairing within this element eliminate NC binding to this site and severely impair RNA packaging into virus-like particles. NMR studies reveal that a recently discovered small-molecule inhibitor of HIV-1 RNA packaging that appears to function by stabilizing the structure of the leader binds directly to the [UUUU]:[GGAG] helix. Our findings suggest a sequential NC binding mechanism for Gag-genome assembly and identify a potential RNA Achilles’ heel to which HIV therapeutics may be targeted.Selective packaging of the HIV-1 genome during virus assembly is mediated by interactions between the dimeric 5ʹ-leader of the unspliced viral RNA and the nucleocapsid (NC) domains of a small number of assembling viral Gag polyproteins. Here, we show that the dimeric 5′-leader contains more than two dozen NC binding sites with affinities ranging from 40 nM to 1.4 μM, and that all high-affinity sites ( Comb (AUG) 18–31. Alth (AUG) 18, 32, 33, (AUG) 18–36. A co (18CES) , which h (18CES) .CES]2 RNA  (gag start site (U5:AUG interactions) leaving residues A345–A356 as a single-stranded unstructured region (344 or 5′-L), which readily forms dimers when incubated under buffer conditions with physiological-like ionic strength  .The 5′-L of the HIV-1 RNA genome includes the 5′-untranslated region and the first few codons of -1NL4-3) A 29). D. Dgag (r-1NL4-3) B 31). I. Igag  see D. All RN2+  were later found to eliminate the nonspecific binding and unfolding  studies from our laboratory showed that the dimeric 5′-L gave rise to a bipartite NC-binding isotherm under PI buffer conditions , 37. Ininfolding , 30. The1 leader , 37. HowKd of 880 ± 30 nM (Kd = 320 ± 30 nM) . A leader construct without the PBS region  binds 23 ± 1 NC with decreased average Kd , suggesting that the PBS region only has two to three weak NC binding sites. Thus, the more than two dozen specific NC binding sites of the dimeric 5′-L are notably heterogeneous in terms of both binding energetics and binding affinities.Fitting of the ITC data using a single-set-of-sites model with the initial “outlier” data points deleted (gray data point due to endothermic binding in  ± 30 nM . This av± 30 nM) , indicatCES, which lacks the TAR-PolyA and PBS elements, is the minimal region of the 5′-L required for packaging , similar to the isotherms and affinities observed for the apical loop of the Ψ-hairpin . These results indicate that the high-affinity (endothermic) binding sites reside within the ΨCES region of the 5′-L.The finding that the TAR-PolyA and PBS regions have very weak NC binding sites is consistent with the previous reports that Ψackaging , 37. NC  fitting . The iniCES]2 structure comprises a central A-helical duplex that connects two tandem three-way junctions   , and the U5:AUG-H1-Ψ three-way junction . The NC binding stoichiometry and affinities of these [ΨCES]2 subfragments were characterized by ITC (Kd = 340 ± 30 nM) similar to that observed for the apical loop of the Ψ-hairpin  and 2.7 ± 0.3 additional exothermic NC binding sites (Kd = 320 ± 180 nM). The exothermic binding sites across different subdomains of [ΨCES]2 exhibit similar affinities with Kd in the range of 300 to 400 nM  , the tand by ITC B–D. The -hairpin . In conto 400 nM . Thus, a3WJ−1. The structure of the junction was previously characterized in the context of a larger RNA (155 nt) comprising half of the symmetric [ΨCES]2 dimer   . The imp2H-edited two-dimensional (2D) nuclear Overhauser effect spectroscopy (NOESY) spectra for AH-, GH-, CH-, or UH-labeled Ψ3WJ−1 RNAs  were assigned using standard methods    . 2H-editterated) B. A311-Hterated) A. Howeveterated) B. The stterated) , possiblterated) .H-labeled Ψ3WJ−1 sample  were substituted by cytosines, thereby converting two G:U base pairs in Ψo Ψ3WJ−1 A. These HP) , and the other with helix-stabilizing 5′-GG and CC-3′ terminating residues at the 5′- and 3′-ends, respectively (ΨHP-stable) . NMR chemical shifts and cross-peak patterns observed for the nonstabilized ΨHP construct were similar to those of corresponding residues in Ψ3WJ−1 , indicating that the structures are similar. Spectra for both constructs exhibited sequential H8/H6-to-H1′ and cross-strand adenosine-H2 to H1′ NOEs for residues 326-AAGGAG-331 and 306-UUUUGA-311  consistent with a helical structure. However, Watson–Crick imino signals for the [UUUU]:[GGAG] element were broadened beyond detection in 1H NMR data obtained for ΨHP but were readily observable in spectra obtained for ΨHP-stable  followed by exothermic binding , whereas the RNA containing helix-stabilizing G–C closing base pairs (ΨHP-stable) exhibited only exothermic NC binding behavior  . Moreover, the Ψ-apical loop and the [UUUU]:[GGAG] element both contain an identical “GGAG” sequence but exhibit distinct binding energetics and affinities  . These ffinities . Thus, iSI Appendix, Fig. S6). However, the packaging efficiency of the U308/309C mutant was substantially diminished compared to that of the wild-type vector (13 ± 2%)  was previously shown to display competitive RNA packaging levels indistinguishable from those of HIV-1 gRNAs . The U3013 ± 2%) E. Thus, 3WJ−1 RNA using 2H-edited NMR. Many 1H NMR signals of NSC exhibited substantial chemical shifts (and some line broadening) when bound to a A2rGrUr-labeled Ψ3WJ−1 sample , suggesting specific interactions between the compound and RNA. Intermolecular NOEs between the RNA ribose protons and the compound were also distinguishable  were barely perturbed upon NSC binding, whereas the H2 signals of adenosines near stem region of the Ψ-hairpin exhibited substantial broadening  was shown to inhibit HIV-1 RNA packaging, apparently by binding to the Ψ-hairpin region of the HIV-1 5′-L and exerting a global stabilizing effect on the RNA structure . NSC wasuishable B. Severaensities A. Two-dioadening C and D. Early efforts to identify HIV-1 RNA packaging determinants identified stretches of nucleotides surrounding (and including) the Ψ-hairpin that, if deleted, resulted in severe HIV-1 packaging defects –25, and The present findings reveal that the conserved [UUUU]:[GGAG] element located at the base of the Ψ-hairpin and adjacent to a tandem three-way junction structure is critical both for high-affinity NC binding and competitive RNA packaging. Each of these two elements in the dimeric 5′-L binds two NC molecules with affinities (∼40 nM) approximately an order of magnitude greater than those of other NC binding sites within the encapsidation-promoting region of the viral leader, and more than 30 times tighter than NC binding sites in other regions of the leader . The absKd ∼100 nM  is a two-component RNA structure, with the “UGCG” tetraloop of SLC bound by the N-terminal zinc finger of NC and the “AUG” residues linking SLA and SLB recognized by the C-terminal zinc finger  (2O (940 μL) was added to yield 1-mL DNA template, which was directly used in the transcription reaction.The DNA templates for the transcription of DIS and Ψ-hairpin derivative RNAs were prepared by annealing a 17-nt T7 promoter sequence  to the corresponding reverse oligonucleotides (IDT): DIS, 5′-mGmGC CCC TCG CCT CTT GCC GTG CGC GCT TCA GCA AGC CGA GTC CTG CCT ATA GTG AGT CGT ATT A-3′; Ψymerase) . BrieflyΔPBS; 5′-mGmGC ACC CAT CTC TCT CCT TCT AGC CTC C-3′ for ΨCES, ΨT−3WJ, Ψ3WJ−1, and Ψ3WJ−1 U308/309C; 5′-mCmAC TAC TTT GAG CAC TCA AGG CAA GC-3′ for TAR-PolyA.The DNA templates for other larger RNA constructs were obtained by standard PCR amplification  using pUC19 plasmids containing the Top17 sequence and the RNA encoding sequences. A common forward amplification primer 80 nt upstream to the T7 promoter sequence was used for all constructs (5′-GGG ATG TGC TGC AAG GCG ATT AAG TTG GG-3′). The reverse amplification primers are 5′-mCmGC ACC CAT CTC TCT CCT TCT AGC CTC C-3′ for 5′-L, 5′-L U308/309C, and 5′-LΔPBS, and ΨCES were described previously . The plasmid for Ψ3WJ−1 was generated using the same overlapping PCR method with 5′-CTG TTG TGT CGA GAG ACG CCA AAA ATT TTG AC-3′ and 5′-TTT TGG CGT CTC TCG ACA CAA CAG ACG-3′ as the two overlapping primers. Plasmid for 5′-L U308/309C was prepared by site-directed mutagenesis  on 5′-L plasmid using forward primer 5′-GGT GAG TAC GCC AAA AAT TCC GAC TAG CGG AGG CTA GAA GG-3′ and reverse primer 5′-CCT TCT AGC CTC CGC TAG TCG GAA TTT TTG GCG TAC TCA CC-3′. Plasmid for Ψ3WJ−1 U308/309C was prepared similarly using Ψ3WJ−1 plasmid and forward primer 5′-GTC GAG AGA CGC CAA AAA TTC CGA CTA GCG GAG GCT AGA AGG-3′ and reverse primer 5′-CCT TCT AGC CTC CGC TAG TCG GAA TTT TTG GCG TCT CTC GAC-3′.The construction of plasmids for 5′-L, 5′-Leviously , 37. The2, 3 to 6 mM NTPs, 2 mM spermidine, 2 mM DTT, 20% (vol/vol) DMSO, 80 mM Tris⋅HCl (pH 9.0), and 0.15 mg of T7 RNA polymerase. The reaction was quenched after a 5-h incubation at 37 °C by addition of an EDTA-urea mixture . The reaction mixture was boiled for 5 min and snap cooled on ice for 5 min prior to addition of glycerol . RNAs were purified by electrophoresis on urea-containing polyacrylamide denaturing gels  at 20 W overnight, visualized by UV shadowing, and eluted using the Elutrap electroelution system (Whatman) at 150 V overnight. The eluted RNAs were concentrated and washed twice with 2 M high-purity NaCl followed by extensive desalting using Amicon Ultra Centrifugal Filter Device (Millipore).RNAs were prepared by in vitro transcription using T7 RNA polymerase (purified in-house) in 7.5- to 30-mL reactions. A 7.5-mL reaction contained ∼0.25 mg of PCR-amplified DNA template or ∼4 nmol of annealed DNA template, 20 mM MgClD, GD, CD, and UD) and partially deuterated (Ur) NTP reagents used for in vitro transcription were purchased from Cambridge Isotope Laboratories (CIL). A2r and Gr were obtained by deuteration of the C8 position of fully protonated ATP and GTP. A volume of 140 µL of triethylamine (TEA) was added to 0.2 g of ATP or GTP dissolved in 8 mL of 99.8% D2O (CIL). One-milliliter aliquots of ATP and GTP were incubated at 60 °C for 5 d and 24 h, respectively. TEA was subsequently removed by lyophilization.Perdeuterated (ANL4-3 NC protein (55 amino acids with the sequence of MQKGN FRNQR KTVKC FNCGK EGHIA KNCRA PRKKG CWKCG KEGHQ MKDCT ERQAN) was transformed into BL21 (DE3) pLysE. The protein was overexpressed and purified as described previously  was loaded into the injection syringe. The calorimetry cell was loaded with 200 µL of RNA (0.6 to 4 μM) in the same buffer as NC. After thermal equilibration at 30 °C and the initial 60-s delay, a single injection of 0.2 μL followed by 19 serial injections of 2 μL were made into the calorimetry cell.ITC experiments were carried out using an ITC200 . A volume of 40 µL of NC (120 to 240 μM) in ITC buffer  on ice to prevent thermal denaturation during electrophoresis.RNA (1 µM) was prepared in PI buffer with 10 mM Tris⋅HCl (pH 7.5), 140 mM KCl, 10 mM NaCl, and 1 mM MgCl1H assignments were obtained from 2D NOESY experiments  using samples in D2O . Imino 1H assignments were obtained from 2D NOESY experiments with jump-return water suppression (T = 10 °C) using samples in 90% H2O/10% D2O. The NSC compound  used for NMR titration experiments was dissolved in DMSO-d6 (CIL) with a final concentration of 20 mM and stored at −20 °C. All NMR data were processed with NMRFx (All NMR data were collected with 600- or 800-MHz Bruker AVANCE spectrometer equipped with cryoprobe. NMR samples were prepared in 10 mM Tris-d11 buffer with an RNA concentration of ∼200 µM in a standard 5-mm NMR tube (500 µL). Nonexchangeable th NMRFx  and analth NMRFx .+ helper NL4-3 GPP contains HIV-1 NL4-3 sequences with env replaced by a puromycin-resistance expression cassette and has been described previously . Products of RPA were quantified by phosphorimaging using standard methods.RNA content of transfected cells and viral particles was analyzed by RNase protection assay (RPA) as previously described . The chiSupplementary File


Severe acute respiratory syndrome coronavirus-2 (SARS-CoV-2), the causative agent of a potentially fatal disease named coronavirus disease 2019 (COVID-19), has raised significant public health concerns globally. To date, the COVID-19 pandemic has caused millions of people to be infected with SARS-CoV-2 worldwide. It has been known since the 2003 SARS epidemic that coronaviruses (CoVs) have large RNA genomes, the replication of which requires an RNA-dependent RNA replication/transcription complex. CoV nonstructural proteins (Nsps) play pivotal roles in the assembly of this complex and associated enzymatic functions in virus genomic replication. Several smaller nonenzymatic Nsps assist with RNA-dependent RNA polymerase function. In this study, we determined the structure of SARS-CoV-2 nonstructural protein 9 (nsp9), an RNA-binding protein that is essential for CoV replication. Its homotetrameric structure with two stable dimeric interfaces provids a structural basis for understanding the mechanisms of RNA-binding protein self-assembly, which may be essential for the regulation of viral RNA replication and transcription. Coronaviridae [Betacoronavirus, which is most closely related to the SARS-like virus previously identified in wild bats [Coronavirus disease 2019 (COVID-19), an acute respiratory distress syndrome caused by severe acute respiratory syndrome coronavirus-2 (SARS-CoV-2), is currently a global pandemic that has been spreading across all continents since late 2019 . Accordiaviridae . Genomicild bats . Human bild bats . Howeverild bats .The genome of SARS-CoVs consists of an approximately 30-kb non-segmented positive-sense RNA sequence with a 5′ untranslated region (UTR), followed by a single open reading frame and a short flanking 3′ UTR . The lifnsp9 gene through a reverse genetics approach was found to prevent the propagation of the virus [As a member of the oligosaccharide/oligonucleotide-binding (OB-fold) superfamily, the RNA-binding protein nsp9 is critical to the RNA-dependent RNA replicase machinery of CoVs  nsp9 is he virus . The seqhe virus , 21–24.To elucidate the molecular mechanisms of SARS-CoV-2 replication, which would aid in the diagnosis, treatment, and prevention of the COVID-19 pandemic, researchers have made significant efforts to characterize the Nsp structures in a short amount of time since the COVID-2019 outbreak . In this1–β7) flanked by an N-terminal extension and a C-terminal α-helix (α1) , avian infectious bronchitis virus (IBV) nsp9 (PDB: 5C94), human coronavirus (hCoV) 229E nsp9 (PDB: 2 J97), porcine delta coronavirus (PDCoV) nsp9 (PDB: 5YM6), and porcine epidemic diarrhea virus (PEDV) nsp9 (PDB: 5HIZ) –24. A suQZ8, avia2  are engaged in the tetrameric structure in two combination modes (helix interface and sheet interface). Interfaces Ia/b (between molecules a and b) and Ic/d (between molecules c and d) share the same combination mode , having a buried area of 626 Å2 and 652 Å2, respectively. Each helix interface involves the conserved C-terminal α-helices (residues 96–109) that are associated in parallel and the N-terminus of nsp9 (residues 1–9)  Fig. a and b. b/c (between molecules b and c) is a unique binding surface located in the center of the SARS-CoV-2 nsp9 tetrameric structure  and three connection loops  from both molecules b and c. The buried area of the Ib/c interface is 1040 Å2, which is the largest buried area compared with those of interfaces Ic/d and Ia/b in the SARS-CoV-2 nsp9 tetramer. There are three major contact regions in the Ib/c interface. The first contact region associates with two LP1 loops (residues 33–42), connecting β2 and β3 from each of the subunits of c and b. Part of the N-terminal tail (residues 2–5) from molecule c is also involved in this contact surface. Two hydrogen bonds and one salt bond are involved in the interface between residues E3 and T35 in molecule c and residue K36 in molecular b  and LP3 loops (residues 91–96) and β5 (residues 64–68) from each of the subunits. The LP2 loop on molecule c comes in direct contact with LP3 on molecule b. An antiparallel β-sheet located at the center of the sheet interface is composed of two β5 strands. In this hydrophobic contact surface, three residues  on molecule c form four hydrogen bonds with residues K92, T67, and Y66 on molecule b, respectively  involved in the nsp9 tetrameric interface. Therefore, we propose that these two dimeric forms are the fundamental units for building the high-order oligomers of nsp9. According to two different research groups that studied nsp9 mutants, the mutation of conserved residues located on the C-terminal α-helix ) or the unexpected appearance of a disulfide on β5 (hCoV-229E nsp9 wild type (PDB: 2 J97)) could significantly change the dimer interface of nsp9 and its ability to bind with DNA or RNA [In this study, we first revealed the horseshoe-like tetrameric structure of nsp9 encoded by the genomic RNA of SARS-CoV-2. This homotetrameric structure comprised two significant styles of dimeric conformation that were found in nsp9 structures published in the last decade , 21. TheA or RNA , 22. In nsp9 gene  was synthesized and cloned into the pET-28a (+)-SUMO vector using BamHI and XhoI sites . The recombinant plasmids, which were confirmed by DNA sequencing , were then used to transform Escherichia coli BL21 (DE3) . Cells were cultured in Luria broth medium at 37 °C until the optical density at 600 nm reached 0.6–0.8. Protein expression was then induced by incubating the cells with 0.5 mM isopropyl β-d-1-thiogalactopyranoside at 16 °C for 18 h. The nsp9 protein was labeled with a His-SUMO tag with a Ulp1 cleavage site, purified by nickel-nitrilotriacetic acid affinity chromatography , and then cleaved with Ulp1 protease. The cleaved and tag-removed protein was loaded onto a Resource S chromatography column  and eluted with a linear gradient of 10 mM to 1 M NaCl. The peak fraction containing the target protein was pooled, then concentrated to 1 mL using Amicon Ultra concentrators , and finally loaded onto a Superdex 200 column  for further purification with a buffer composed of 20 mM Tris (pH 8.0), 150 mM NaCl, 1 mM dithiothreitol, and 10% glycerol (v/v). The purity of the proteins was greater than 95%, as confirmed by sodium dodecyl sulfate polyacrylamide gel electrophoresis 2SO4 after growth for 2 days at 20 °C 2SO4, and the mixture was then frozen and stored in liquid nitrogen for future data collection.Using the hanging-drop vapor diffusion method, the nsp9 protein was quantified using absorbance readings at A280 nm  and concentrated to 17 mg/mL for the crystallization trials, by mixing 1 μL of protein with 1 μL of reservoir solution at 20 °C. Commercial crystallization kits  were used to obtain the initial crystallization conditions. The protein crystals were obtained in a solution containing 100 mM citric acid (pH 3.0), 200 mM potassium sodium tartrate tetrahydrate, and 2.0 M  at a wavelength of 0.97930 Å. Data were processed and scaled using the HKL2000 package , in a solution containing 10 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (pH 8.0), 50 mM KCl, 1 mM ethylenediaminetetraacetic acid (EDTA), 0.05% Triton-X-100, and 5% glycerol, for 30 min at 4 °C. Thereafter, a 10× loading buffer (250 mM Tris-HCl (pH 7.9) and 40% glycerol) was added to the mixture. Then, samples were electrophoresed on 6.5% non-denaturing Tris–borate–EDTA (TBE) polyacrylamide gels for 40 min at a voltage of 100 V, and the results were determined with a ChemiDoc Touch Imaging system .The structure factors and atomic coordinates have been deposited in the Protein Data Bank under the PDB ID codes 7BWQ.Additional file 1: Figure S1. The OB-fold cluster of SARS-CoV-2 nsp9. (a) The different layers are colored orange, light blue, and pale green, respectively, and protomers in every layer are depicted in cartoon representations and labeled with uppercase letters. (b) Molecules d’, c’, asym, and bsym in the stable tetramer are colored in bright orange, pale green, light blue, and orange, respectively, and depicted in cartoon and surface representations.Additional file 2: Figure S2. Purification and X-ray data collection of SARS-CoV-2 nsp9. (a) The chromatography of the purification of SARS-CoV-1 nsp9 with a Superdex200™ 10/300 column. (b) SDS-PAGE analysis result of SARS-CoV-2 nsp9. M is the protein marker. (c) The primary crystal state of SARS-CoV-2 nsp9 in the commercial crystallization kit. (d) The crystal for X-ray data collection of SARS-CoV-2 nsp9 after crystallization condition optimized. (e) X-ray diffraction pattern for structure analysis (left) and X-ray diffraction point atlas of SARS-CoV-2 nsp9 (right).Additional file 3: Table S1. PDBePISA calculation of interface parameters of the quaternary structure of SARS-CoV-2 nsp9.


The T‐box riboswitches are widespread bacterial noncoding RNAs that directly bind specific tRNAs, sense aminoacylation on bound tRNAs, and switch conformations to control amino‐acid metabolism and to maintain nutritional homeostasis. The core mechanisms of tRNA recognition, amino acid sensing, and conformational switching by the T‐boxes have been recently elucidated, providing a wealth of new insights into multivalent and multimodal RNA–RNA interactions. This review dissects the structures and tRNA‐recognition mechanisms by the Stem I, Stem II, and Discriminator domains, which collectively compose the T‐box riboswitches. It further compares and contrasts the two classes of T‐boxes that regulate transcription and translation, respectively, and integrates recent findings to derive general themes, trends, and insights into complex RNA–RNA interactions. Specifically, the T‐box paradigm reveals that noncoding RNAs can interact with each other through multiple coordinated contacts, concatenation of stacked helices, and mutually induced fit. Numerous tertiary contacts, especially those emanating from strings of single‐stranded purines, act in concert to reinforce long‐range base‐pairing and stacking interactions. These coordinated, mixed‐mode contacts allow the T‐box RNA to sterically sense aminoacylation on the tRNA using a bipartite steric sieve, and to couple this readout to a conformational switch mediated by tRNA‐T‐box stacking. Together, the insights gleaned from the T‐box riboswitches inform investigations into other complex RNA structures and assemblies, development of T‐box‐targeted antimicrobials, and may inspire design and engineering of novel RNA sensors, regulators, and interfaces.RNA Structure and Dynamics > RNA Structure, Dynamics and ChemistryRegulatory RNAs/RNAi/Riboswitches > Regulatory RNAsRegulatory RNAs/RNAi/Riboswitches > RiboswitchesThis article is categorized under: Recent elucidation of the structures and mechanisms of bacterial T‐box riboswitches provides insights into how structured noncoding RNAs interact with each other and regulate gene expression through multivalent contacts that go much beyond base pairing. Through4.2N. farcinica and M. tuberculosis ileS structures show that the Stem IIA/B pseudoknot makes no direct contact with the tRNA  are in fact structural components of the Stem IIA/B pseudoknot facing Stems I and II, forming a key feature required for tRNA binding (Suddala & Zhang, The structural and functional elucidation of Stem II domain resolves several long‐standing mysteries in the T‐box mechanism. First, it explains why the glycine‐specific Taken together, specific recognition of the tRNA identity by the Stem I and II domains sets the stage for the T‐box discriminator domain to probe the tRNA 3′‐end to check for aminoacylation, and to couple this readout with an RNA conformational change that governs downstream gene expression.5The most essential and unique feature of the T‐box mechanism is the ability to sense the presence or absence of an esterified amino acid on the tRNA 3′‐end and regulate downstream gene expression accordingly. This is no easy task and unprecedented for a small RNA‐only device, which comprises only about 60 nts. This function is executed by a highly compact structure termed the T‐box discriminator, and was only recently defined based on its structural and functional elucidation (Li et al., 5.1Pioneering genetic and biochemical work established that in addition to the three codon–anticodon base pairs from Stem I, a second set of base‐pairing interactions form between the tRNA 3′‐NCCA terminus and the so‐called T‐box bulge Table . It was B. subtilis glyQ T‐box that the molecular volume of the tRNA 3′ group is the best predictor of the tRNA‐mediated transcription readthrough (Zhang & Ferré‐D'Amaré, cis diols present on the RNA 3′‐end, does not distinguish among the amino acids and reject them blanketly. It also rejects other chemical moieties that are not amino acids, such as azido and phosphate groups, suggesting a general steric sieve that is intolerant of any significant bulk larger than ~60 Å3 at the tRNA 3′‐end (Zhang & Ferré‐D'Amaré, With the ability to sense tRNA aminoacylation confirmed, the crux of the problem became how an RNA can read a minute chemical change on another RNA. Using a series of tRNA variants that carry diverse chemical adducts to the ribose of tA76, it was demonstrated using 5.2−1, it is necessary to guard against both types, to ensure efficacious sensing of tRNA aminoacylation. In conclusion, the new discriminator‐tRNA complex structures not only validated the proposed steric sensing mechanism based on biochemical analyses (Zhang & Ferré‐D'Amaré, As is with any steric occlusion mechanism, a snug binding pocket must exist on the T‐box discriminator that fully encompasses the tRNA 3′‐end. This steric pocket is recently visualized by two highly similar co‐crystal structures—one transcriptional and the other translational—and further confirmed by a cryo‐EM structure, covering three different species (Battaglia et al., 5.3After tRNA aminoacylation is sterically sensed, how is the readout communicated and coupled to genetic switching? In other words, how does the uncharged tRNA specifically stabilize the “ON” conformation of the discriminator? The key lies in an intermolecular coaxial stacking interaction between the tRNA 3′‐end and Helix A1 of the T‐box discriminator. Intramolecular coaxial stacking of adjacent dsRNA helices within a complex RNA molecule is principally responsible for shaping the overall architecture of structured RNAs (Butcher & Pyle, Three independently solved structures eventually validated this model and directly visualized the functionally critical tRNA–Helix A1 stacking interaction (Battaglia et al., B. subtilis full‐length T‐box complex detected an “ultrastable” complex with the half‐life of about 1 hr, revealing the unusual stability of this multivalent complex (Suddala et al., Remarkably, the tRNA‐Helix A1 coaxial stack extends further upstream into the tRNA acceptor and T stems Table , creatinTaken together, the original tRNA‐T‐box stacking concept is supported by biophysical characterizations using fluorescence probing and calorimetry, biochemical characterizations employing in vitro termination‐readthrough analyses, and complemented by recent high‐resolution structural data. These findings suggest a potentially general mechanism of gene‐expression control mediated by intermolecular RNA–RNA stacking, and provide a proof‐of‐principle that one structured RNA can act as a trans‐activator for another RNA conformational switch, through a combination of base‐pairing, stacking, and tertiary interactions. Based on this concept, RNA therapeutics can be potentially designed to manipulate conformations of bistable RNA elements to control gene expression, splicing, microRNA targeting, and so on.6A number of differences exist between the two major classes of T‐boxes that regulate transcription and those modulating translation, both in their regulatory behavior as well as sequence and structure Figure . The nex6.1One predominant difference between the two types of T‐boxes is the distinct kinetic behavior of their regulatory mechanisms. Transcriptional T‐boxes are “one‐off” switches that make a single genetic decision within a short time window which either permits or forbids an elongating RNAP from traversing a “checkpoint” region in DNA. This region encodes a hairpin‐dependent, intrinsic transcription terminator—a strong RNA hairpin followed by a track of uridines—which guards the entrance into the downstream open reading frame (Grundy & Henkin, In stark contrast, translational T‐boxes are bistable RNAs that are chemically homogenous but conformationally distinct. At least two conformational states are in rapid equilibrium, enabling the RNA to continuously monitor the charging levels of specific tRNAs and change conformations accordingly. Here, the “ON” and “OFF” states do interconvert and do not correspond to chemically distinct species, but rather different conformational states of the same RNA. By exposing or sequestering the Shine–Dalgarno sequence to which ribosomal RNA binds, the “ON” and “OFF” conformational states control translation initiation. Regulation does not require active transcription as do the transcriptional T‐boxes. These decidedly different modes of tRNA interaction and structural switching between the two T‐box classes presumably drove the adoption of distinctive sequence and structural features discussed below.6.2Staphylococci glyQ T‐boxes (Apostolidi et al., A principal difference in the architecture of transcriptional and translational T‐boxes is the length and variability of Stem I. Transcriptional T‐boxes generally require long Stem I's featuring the IDTM motif that binds the tRNA elbow. Mutations that disable the IDTM significantly compromise T‐box function in vitro and in vivo (Suddala et al., Importantly, the two classes of T‐boxes share a highly congruent core structure which captures and probes the tRNA 3′‐end and actuates switching via tRNA–mRNA stacking (Battaglia et al., 7The recent flourish of genetic, biochemical, single‐molecule, and structural analyses into the T‐box riboswitches have shined a timely light on understanding multivalent and multimodal RNA–RNA interactions in general. Three chief conceptual advances gleaned from the T‐box paradigm are discussed below.First, RNA–RNA interactions can effectively leverage multivalency. The archetypal T‐box system features three RNA–RNA interfaces that are interestingly roughly equidistant from each other at ~60 Å apart Figure . These din crystallo formation of pseudo‐infinite helices (Shoffner, Wang, Podell, Cech, & Guo, Second, RNA–RNA interactions can employ multi‐segment, concatenated intermolecular stacking. It is well recognized that intramolecular coaxial stacking between dsRNA helices is a chief sculptor for overall RNA architectures (Dussault, Dube, Jacques, Grondin, & Lafontaine, Third, RNA–RNA interactions can utilize mutually induced fit. Large RNAs are dynamic, flexible entities and can possesses multiple domains. Just like protein–protein interactions, major conformational changes in both the T‐box and the tRNA accompany and enable their binding. The Stem I hinge near the C‐loop pivots in order to accommodate the flexing of the tRNA t26•t44 hinge. In translational T‐boxes to which tRNAs reversibly bind and dissociate from, the discriminator domain most likely has to reorganize its fold upon tRNA binding and dissociation. In particular, the string of purines that flank Stem III presumably can only bind after its binding site forms, which is the minor groove of the tRNA‐T‐box heteroduplex. Upon tRNA departure, the loss of the positioning contacts will likely lead to dissociation of the Stem III purine string and partial collapse of the discriminator structure. The drastic structural differences between an isolated T‐box antiterminator domain and its tRNA complex lends support to the notion of induced fit by tRNA (Gerdeman, Henkin, & Hines, These critical themes and insights gathered from the T‐box paradigm are potentially applicable in a wide range of large noncoding RNAs, RNA condensates, and RNP assemblies, and are expected to usher in a new wave of structural and mechanistic elucidations of complex noncoding RNAs.8B. subtilis glyQS T‐box remains the only T‐box that has been successfully reconstituted in vitro with defined components. Intriguingly, the B. subtilis thrS T‐box gained function in vitro upon the addition of a partially purified protein fraction, suggesting that there could be additional components in the T‐box regulon (Putzer et al., Despite the recent unraveling of the core T‐box mechanism (Weaver & Serganov, First, it remains unclear to what extent the T‐box sequence and structure have been gradually sculpted by the emergence of numerous post‐transcriptional modifications on their tRNA ligands through their co‐evolution. T‐box riboswitches are ancient gene‐regulatory elements that might have existed in the RNA world considering its functional independence from proteins (Zhang & Ferré‐D'Amaré, Second, we lack a detailed kinetic picture comprising the sequence and timing of a series of discrete binding events between the tRNA and T‐box RNA and of the conformational switching that follows. The intimate, short‐range Stem II S‐turn interactions with the codon–anticodon duplex, despite engaging numerous hydrogen bonds, lack base complementarity and thus the addressing power to locate to its target site. Thus, the S‐turn likely can only serve as a secondary contact and requires spatial guidance by the Stem I, K‐turn and pseudoknot. Considering the 5′‐to‐3′ polarity of transcription for the major class of transcriptionally acting T‐boxes (Zhang & Landick, Third, there exist a large variety of lineage‐specific alterations to the core T‐box architecture. These features likely evolved in adaption to the diverse environmental conditions, nutritional profiles and regulatory needs of the microorganisms. There are fully and partially tandem T‐boxes whose modes of action remain unknown. There are curious insertions in every domain of the T‐box with unknown structure and function. The less conserved Stem II domain further harbors remarkable diversity in sequence and structure. These insertions may engage novel contacts with other regions of the tRNA. Given the large surface area and complex scaffold of tRNAs as compared to small‐molecule metabolites, it is not surprising that through evolution, the primordial T‐boxes have diverged and accrued a large collection of variations. Additionally or alternatively, some of these lineage‐specific elements may have evolved to permit crosstalk and coordination with other forms of RNA or ribonucleoprotein‐mediated gene regulation. For instance, a tandem arrangement of a T‐box and a ppGpp riboswitch was reported to form a Boolean “AND” gate (Sherlock, Sudarsan, Stav, & Breaker, Finally, the recent elucidation of the core T‐box mechanisms has informed and continues to inspire rational design and engineering of RNA devices that can recognize the 3D structure of other RNAs or proteins, sense minute chemical modifications on them, catalyze desired chemical transformations on specific RNA targets, or assemble into programmed suprastructures using joints and interfaces ported from the T‐box paradigm. T‐boxes can serve as effective tools in synthetic biology thanks to their unique ability to decode specific tRNAs in the RNA form without translation. An outstanding example is the newly minted T‐box ribozyme named “Tx2.1”, which harnessed the tRNA‐binding specificity of the T‐box RNA to allow its flexizyme‐like moiety to aminoacylate specific tRNAs like their protein counterparts—the aminoacyl‐tRNA synthetases (aaRS; Ishida, Terasaka, Katoh, & Suga, The author declares no conflicts of interest.Structure and mechanism of the T‐box riboswitches