Biochemical and Biophysical Research Communications 389 (2009) 616–621

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Biochemical and Biophysical Research Communications

journal homepage: www.elsevier .com/locate /ybbrc

Fluorescence probing of T box antiterminator RNA: Insights into riboswitchdiscernment of the tRNA discriminator base

John A. Means, Crystal M. Simson, Shu Zhou, Aaron A. Rachford, Jeffrey J. Rack, Jennifer V. Hines *

Department of Chemistry and Biochemistry, Ohio University, Athens, OH 45701, USA

a r t i c l e i n f o a b s t r a c t

Article history:Received 8 September 2009Available online 13 September 2009

Keywords:RNARiboswitchT boxTranscription2-AminopurineFluorescence

0006-291X/$ - see front matter � 2009 Elsevier Inc. Adoi:10.1016/j.bbrc.2009.09.037

* Corresponding author. Fax: +1 740 593 0148.E-mail address: hinesj@ohio.edu (J.V. Hines).

The T box transcription antitermination riboswitch is one of the main regulatory mechanisms utilized byGram-positive bacteria to regulate genes that are involved in amino acid metabolism. The details of theantitermination event, including the role that Mg2+ plays, in this riboswitch have not been completelyelucidated. In these studies, details of the antitermination event were investigated utilizing 2-aminopu-rine to monitor structural changes of a model antiterminator RNA when it was bound to model tRNA.Based on the results of these fluorescence studies, the model tRNA binds the model antiterminatorRNA via an induced-fit. This binding is enhanced by the presence of Mg2+, facilitating the complete basepairing of the model tRNA acceptor end with the complementary bases in the model antiterminatorbulge.

� 2009 Elsevier Inc. All rights reserved.

Introduction

The T box riboswitch is a transcriptional control mechanismfound primarily in Gram-positive bacteria [1–3], and its prevalenceindicates that it is one of the main regulatory mechanisms forgenes involved in amino acid metabolism [4]. The 50 untranslatedleader region (50-UTR) of the T box genes is composed of a complexset of conserved primary sequence and secondary structural ele-ments that allow the nascent mRNA to sense the charging ratioof the gene’s cognate tRNA [5,6]. Control of T box genes hingeson the formation of one of two mutually-exclusive, competingstructural elements—the terminator and the antiterminator [1–3]. The uncharged cognate tRNA is capable of factor-independentbinding with the mRNA via a two-point interaction: the tRNA anti-codon with the mRNA specifier sequence and the tRNA acceptorend with the complementary bases in the antiterminator(Fig. 1A) [1,5,7–9]. The stabilization of the antiterminator throughthis interaction allows transcription of the gene, while the forma-tion of the competing terminator (in the absence of uncharged,cognate tRNA) results in transcription termination. The specificityof the interaction between the tRNA and mRNA is dictated by theco-variation of the tRNA anticodon and the mRNA specifiersequence [1] and by the co-variation of the tRNA discriminatorbase and the mRNA variable base, which is located in the bulgeof the antiterminator element [5].

Interesting facets of this unique interaction between the 50-UTRof the mRNA and the tRNA have been elucidated via in vitro

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biochemical studies. In one study involving the Bacillus subtilis gly-QS 50-UTR, Mg2+ was implicated as an important requirement forefficient in vitro antitermination [9,10]. Near maximal tRNA bind-ing occurs with 5 mM Mg2+, but maximal antitermination is notachieved until 15 mM Mg2+. In another in vitro study utilizing thissame model system, it was found that the tRNA is able to be com-petitively displaced prior to the formation of the antiterminator.However, once the tRNA is bound to the mRNA at both points ofcontact, the tRNA is resistant to competitive displacement [10].

The 2-aminopurine purine base analog (AP) provides local nu-cleic acid structural information via steady-state [11–15] andtime-resolved [16–19] fluorescence experiments. The fluorophoresenses local stacking and polarity changes, yielding changes inemission intensity and/or emission wavelength [20–22]. In addi-tion, AP is capable of base pairing with uracil in a manner that issimilar to that of adenine [23–25]. These characteristics of AP haveallowed the substitution of the fluorophore for a variety of nucleo-tides, especially guanine and adenine, to monitor nucleic aciddynamics and interactions with minimal structural perturbationto double-stranded motifs and a variety of single-stranded RNAmotifs [11,13–18,21,24–27].

Past studies from this lab have raised some interesting hypoth-eses regarding the interaction between the tRNA acceptor end andthe antiterminator. The NMR solution structure of an antitermina-tor model RNA, AM1A [28], revealed structural flexibility in theantiterminator that was hypothesized to allow for tRNA bindingvia an induced-fit and/or tertiary structure capture. A more recentstudy identified a preferred binding site for Mg2+ along helix A1 inAM1A [29]. From this work, it was hypothesized that the Mg2+

binding at this site might facilitate tRNA binding by reducing

Fig. 1. (A) Schematic of T box gene mRNA leader region with cognate uncharged tRNA. The tRNA binds to the leader region via two points of contact: (1) tRNA anticodon withthe specifier sequence in Stem 1 of the leader region [1], and (2) tRNA acceptor end with the complementary bases in the 50 end of the antiterminator bulge [5]. (A1 and A2refer to the helical regions in the antiterminator). Antiterminator and tRNA model RNAs. (B) 2-AP-AM1A; (C) 9-AP-AM1A [30]; (D) 10-AP-AM1A; (E) 13-AP-AM1A; (F) tRNA-UCCA [28]; (G) mh-UCCA [28] and mh-ACCA [28].

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electrostatic repulsion. In the current work, AP was utilized toprobe structural changes in T box antiterminator model RNA uponits binding with model tRNA. The results of these studies will pro-vide evidence in support of the two hypotheses presented above. Inaddition, while much work has been done to increase the generalunderstanding of the nature of AP in DNA dynamics, these studieswill contribute to the understanding of the nature of AP in RNAdynamics.

Materials and methods

Model RNAs. Local environmental changes within the structureof the antiterminator model RNA, AM1A, were monitored usingthe 2-aminopurine fluorescent purine base analog (AP). TheAM1A model RNA motif has been demonstrated, both in vitroand in vivo, to be structurally and functionally relevant [28]. Fourdifferent AP-labeled AM1As (Fig. 1B–E) were designed to monitorchanges in the A1 helix, the bulge, and the hinge region. In theA1 helix, AP was substituted for the adenine at position 2 (2-AP-AM1A). For probing the bulge, AP was substituted for the adeninesat positions 9 and 10 (9-AP-AM1A [26,30] and 10-AP-AM1A,respectively). Finally, AP was substituted for the guanine at posi-tion 13 in the hinge region (13-AP-AM1A). Purified, fluores-cently-labeled RNAs were obtained from Dharmacon Research, Inc.

An unlabeled model tRNA [28] (tRNA-UCCA, Fig. 1F) and twounlabeled tRNA acceptor stem/end microhelix model RNAs [28](mh-UCCA and mh-ACCA, Fig. 1G) were bound with the AP-AM1A RNAs in titration and fluorescence decay experiments. Themh-UCCA and tRNA-UCCA have an acceptor end sequence that isfully complementary to the four bases at the 50 end of the AM1Abulge. The mh-ACCA model is the single base mismatch analog,in which the discriminator base is not complementary to theAM1A variable base. All of these model tRNAs, whose functionalrelevance were previously demonstrated [26,28], were synthesizedvia in vitro transcription with T7 RNA polymerase [31–33],

followed by purification with denaturing gel electrophoresis andelectroelution.

All RNAs were dialyzed into 10 mM sodium phosphate pH 6.5,0.01 mM EDTA and were renatured prior to use. In addition, allRNA stock solutions were stored in standard microcentrifugetubes, as opposed to low-binding microcentrifuge tubes. Duringthis investigation, it was discovered that previously reported preli-minary results [26] were contaminated by trace impurities that re-sulted from the use of low-binding microcentrifuge tubes for RNAstock solution storage (data not shown). Due to the large amount ofthe unlabeled microhelix RNAs that was necessary for those previ-ously reported fluorescence titration studies, an emission band at394 nm was observed upon excitation of these trace impuritiesat 310 nm. The absence of the trace impurities in the RNA stocksolutions for the current studies was verified by obtaining the fluo-rescence emission profiles of all unlabeled RNAs (data not shown).In light of the results from the current work, the previous traceimpurities did not affect the binding of the model antiterminatorand model tRNAs in the previous work.

Steady-state fluorescence. The steady-state fluorescence spectrawere obtained by titrating 400 lL of 100 nM AP-RNA (50 mMsodium phosphate pH 6.5, 0.01 mM EDTA, 50 mM NaCl, 15 mMor 5 mM MgCl2) with 2 lL aliquots (up to 20 lL total) of the modeltRNA or either of the microhelix model tRNAs. The experimentswere performed with a JY Horiba Spex Fluoromax-3, which wasequipped with a temperature-controlled sample holder that wasset to 20 �C. The labeled RNAs were excited at 310 nm (5 nm slitwidth), and the fluorescence emission was monitored over therange of 330–600 nm (5 nm slit width). Spectra were obtainedfor the unbound, labeled RNAs and after each successive additionof the model tRNAs. The buffer background was subtracted fromeach spectrum to remove the water Raman signal at 347 nm.

Time-resolved fluorescence. Fluorescence decay spectra wereobtained with a PTI LaserStrobe spectrofluorometer that wasequipped with a PTI GL-3300 nitrogen laser as its light source.

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The 2-aminopurine fluorophore was excited at 337 nm, utilizing afrequency of 10 Hz and 5 laser shots per time point. Fluorescenceemission decays were obtained at 370 nm for 1 lM free 2-amino-purine deoxyribonucleoside (APdNS) and at both 370 nm and420 nm for 1 lM 9-AP-AM1A by itself and a 99% bound complexof 9-AP-AM1A and mh-UCCA. The concentration of mh-UCCA thatwas needed to achieve the 99% (1521 lM) bound state was calcu-lated based on the previously reported Kd of 16 lM for 50-Fl-AM1Aand mh-UCCA [26]. All RNA solutions had a volume of 400 lL andcontained 50 mM sodium phosphate pH 6.5, 0.01 mM EDTA,50 mM NaCl, 15 mM MgCl2. The decay spectra were collected withstart and end delays of 50 and 100 ns, respectively. The data pointswere obtained with an integration time of 1000 ls and were anaverage of 7 reads over 200 channels. The excitation and emissionslit widths were adjusted to maximize the intensity of the 9-AP-AM1A fluorescence intensity at 370 nm. The instrument responsefunction was collected at 337 nm, utilizing an empty cuvette anda neutral density filter in the emission path.

Decays were deconvoluted with the TimeMaster software (PTI,Inc.), and the goodness of fit was based on the reduced {2 andthe randomness of the autocorrelation function of the weightedresiduals. The time-resolved fluorescence spectra were fit to thefollowing decay function:

IðtÞ ¼XN

i¼1

aie�t=si

where I(t) is the fluorescence intensity as a function of time, ai is theamplitude for each component, t is the time, si is the fluorescencelifetime of each component, and N is the number of components.In order to determine the minimum number of components thatwere required to fit each of the decay spectra, each spectrum wasfit utilizing N = 1, 2, 3, and 4.

Results

Steady-state fluorescence

Addition of mh-UCCA in the presence of 15 mM MgCl2 led tostrong fluorescence enhancements for the antiterminator modelRNAs 2-AP-AM1A, 10-AP-AM1A, and 13-AP-AM1A (Fig. 2A, E, andG). An enhancement of the fluorescence intensity likely indicates adestabilization of the stacking around the fluorophore, since stack-ing quenches fluorescence [34–36]. When each of these AP-AM1Aswas titrated with mh-ACCA, similar results were observed; however,the intensity enhancements were less pronounced (Fig. 2B, F, and H).The titrations of 2-AP-AM1A and 10-AP-AM1A with either of themicrohelix RNAs yielded no significant emission wavelength shift,but a 6 nm bathochromic shift was observed when 13-AP-AM1Awas titrated with either mh-UCCA or mh-ACCA.

The titration experiments involving 9-AP-AM1A were intrigu-ingly different from the other AP-labeled antiterminator modelRNAs. When 9-AP-AM1A was titrated with mh-UCCA at 15 mMMgCl2, no net intensity enhancement was observed (Fig. 2C). In-stead, the increasing concentrations of mh-UCCA induced a 12 nmbathochromic shift of the emission wavelength and a broadeningof the emission peak. Comparable results were observed when9-AP-AM1A was titrated with tRNA-UCCA in the presence of15 mM MgCl2. Namely, no net intensity enhancement, a broadeningof the emission peak, and a 6 nm bathochromic shift of the emissionwavelength were observed (Fig. 2K). As with the other three AP-AM1A titration experiments, titration of 9-AP-AM1A with the mis-matched mh-ACCA at 15 mM MgCl2 (Fig. 2D) yielded effects thatwere similar to those of mh-UCCA, but were less pronounced.

The microhelix titrations of 9-AP-AM1A were also performed at5 mM MgCl2. In contrast to all the spectra with 15 mM MgCl2, the

spectra from mh-UCCA and mh-ACCA titrations of 9-AP-AM1A at5 mM MgCl2 exhibited similar intensity enhancements for mh-UCCAcompared to enhancements for mh-ACCA, while there was no shift ofthe emission wavelength induced by either microhelix (Fig. 2I–J).

Time-resolved fluorescence

Time-resolved fluorescence measurements were employed tobetter understand the nature of the peak broadening and the12 nm bathochromic shift of the 9-AP-AM1A emission at 15 mMMgCl2 with mh-UCCA. The time-resolved fluorescence data aresummarized in Table 1 and the decay spectra can be found in Sup-plementary data. The decay spectra for APdNS were best fit with amonoexponential function, yielding a decay lifetime of 8.4 ns. Thisis similar to previously-reported decay lifetimes for various free APspecies [17,37–39]. The spectrum for 9-AP-AM1A was best fit witha biexponential function at 370 nm. For 9-AP-AM1A, the shorter ofthe two decay lifetimes was in the range of 2 ns, while the longer ofthe two lifetimes was in the range of 7.6–9.5 ns. At 420 nm, the de-cay profile of the 99% complex was best fit with a monoexponentialfunction, and that single lifetime (2.7 ns) was comparable to theshorter lifetime of the 9-AP-AM1A at 370 nm. The decay lifetimesthat corresponded to 9-AP-AM1A at 420 nm, as well as for the99% complex at 370 nm, were disregarded due to the potentiallyinaccurate nature of the data that resulted from their very lowfluorescence intensities.

Discussion

Induced-fit binding

For each AP-labeled antiterminator model RNA, greater fluores-cence enhancements or kmax shifts were observed when titratedwith mh-UCCA, compared to titration with the mismatched mh-ACCA. These results indicate that base pairing of the tRNA discrim-inator and antiterminator variable bases induces a structuralchange throughout the antiterminator. The most dramatic differ-ences were with 2-AP-AM1A and 9-AP-AM1A. In the case of 9-AP-AM1A, structural studies of uncomplexed AM1A indicated thatA9 is stacked in a hydrophobic pocket with the Watson–Crick facenot readily accessible for base pairing [40]. The more significantkmax shift in 9-AP-AM1A that was induced by the matched discrim-inator base (mh-UCCA), as compared with the shift induced by themismatched discriminator base (mh-ACCA), is consistent with astructural rearrangement involving A9 (the variable base). Thisrearrangement can be correlated to the identity of the discrimina-tor base and may be necessary to accommodate the base pairing ofthe variable base with the model tRNA discriminator base. In thecase of 2-AP-AM1A, previous studies have shown that the phyloge-netically-conserved 50-side of helix A1 is a diffuse Mg2+ binding sitethat may facilitate tRNA binding [29]. The more significantenhancement of 2-AP-AM1A fluorescence in the presence of thematched discriminator base, as opposed to the mismatched dis-criminator base, and the data indicating that the discriminatorbase-variable base pair formation in 9-AP-AM1A is facilitated byhigher Mg2+ concentrations are both consistent with a structuralchange occurring in the region of the second nucleotide in the dif-fuse Mg2+ binding site of helix A1 to facilitate tRNA binding.

Mg2+ facilitation of binding

The data indicate that Mg2+ likely plays a role in facilitatingcomplete base pairing of model tRNA with AM1A (including thediscriminator base-variable base pairing). The titration of 9-AP-AM1A with mh-UCCA at 5 mM MgCl2 led to a simple fluorescenceintensity enhancement. This is in marked contrast to the batho-

Fig. 2. Fluorescence spectra for titrations of AP-AM1A with tRNA model RNAs in 50 mM sodium phosphate pH 6.5, 0.01 mM EDTA, 50 mM NaCl, 15 mM MgCl2 (except wherenoted). (A) 2-AP-AM1A with mh-UCCA, (B) 2-AP-AM1A with mh-ACCA, and (C) 9-AP-AM1A with mh-UCCA; (D) 9-AP-AM1A with mh-ACCA; (E) 10-AP-AM1A with mh-UCCA;(F) 10-AP-AM1A with mh-ACCA; (G) 13-AP-AM1A with mh-UCCA; (H) 13-AP-AM1A with mh-ACCA; (I) 9-AP-AM1A with mh-UCCA in 5 mM MgCl2; (J) 9-AP-AM1A with mh-ACCA in 5 mM MgCl2; and (K) 9-AP-AM1A with tRNA-UCCA. With the exception of 13-AP-AM1A with mh-ACCA, all observed enhancements were more significant than therespective buffer control enhancements.

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Table 1Fluorescence decay lifetimes and amplitudes for free 2-aminopurine deoxyribonucleoside (APdNS; at 370 nm), 9-AP-AM1A (at 370 and 420 nm) and 99% 9-AP-AM1A/mh-UCCAcomplex (at 370 and 420 nm).

Sample a1 s1 (ns) a2 s2 (ns) {2

APdNS (370 nm) — — 0.028 ± 0.0003 8.4 ± 0.1 1.049-AP-AM1A (370 nm) 0.014 ± 0.001 2.1 ± 0.3 0.003 ± 0.001 9.5 ± 1.7 1.299-AP-AM1A (420 nm) naa naa naa naa naa

99% Complex (370 nm) naa naa naa naa naa

99% Complex (420 nm) 0.007 ± 0.001 2.7 ± 0.3 — — 0.94

a Fluorescence decays were measured at 370 nm and 420 nm for each sample and were analyzed individually. Due to very low fluorescence intensities, the data for 9-AP-AM1A at 420 nm and the data for 99% Complex at 370 nm were not utilized, see text.

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chromic shift of the emission band observed for the same titrationat 15 mM MgCl2. A proposed model of these different bindingmodes is shown in Fig. 3.

The results of the steady-state fluorescence titration studies at5 mM MgCl2 may arise from incomplete binding of the mh-UCCAwith 9-AP-AM1A (II in Fig. 3). This is supported by the steady-statefluorescence data comparing the mh-UCCA-induced emission ef-fects with the mh-ACCA-induced emission effects. The fact thatthe mh-ACCA-induced effects were not dissimilar from the mh-UCCA-induced effects likely indicates incomplete binding, as op-posed to a lack of binding.

The steady-state fluorescence data and the time-resolved fluo-rescence data at 15 mM MgCl2 indicate that mh-UCCA is possiblybinding AM1A differently than it does at 5 mM MgCl2. The struc-tural change that is likely induced throughout the antiterminator,as was intimated from the steady-state fluorescence data involvingthe four different AP-AM1A models, may be indicative of a morecomplete binding event than is observed at 5 mM MgCl2 (III inFig. 3). The apparent peak broadening and bathochromic shift ofthe emission wavelength that was observed for the titration of 9-AP-AM1A with mh-UCCA at 15 mM MgCl2 more closely resembledthe appearance of a long wavelength emission band (�420 nm),coupled with the disappearance of the typical AP emission bandat 370 nm. The fluorescence decay profile differences for the emis-sion bands at 370 nm (9-AP-AM1A) and 420 nm (99% complex)support the hypothesis that the emission bands are likely arisingfrom two different emitting conformational species.

Fig. 3. Proposed binding model for formation of tRNA–antiterminator RNA com-plex. In the presence of low concentrations of Mg2+, unbound antiterminator RNA (I)binds the 50-CCA-30 acceptor end nucleotides of tRNA to form II. In the presence ofhigh concentrations of Mg2+, unbound antiterminator RNA (I) binds the entire 50-UCCA-30 acceptor end nucleotides of tRNA to form III.

Peak broadening and long-wavelength shifts have been observedpreviously in DNA [34,41–43]. From recent studies with AP-labeledsingle-stranded and duplex DNA, the emission at the longer wave-length was proposed to result from the formation of a highly-stackednucleodimer between AP and its neighboring base [20]. A similarconclusion was reached in base flipping studies involving the EcoKIand the M.TaqI methyltransferases [20,43,44]. In these studies, thelonger wavelength fluorescence of AP was correlated with the stack-ing of the extrahelical fluorophore with an aromatic sidechain with-in the enzyme. In light of these other studies, the long wavelengthemission band that arises from the titration of 9-AP-AM1A withmh-UCCA at 15 mM MgCl2 could indicate that AP is assuming ahighly-stacked arrangement in the complex.

Conclusions

These studies provide insight into the mechanistic details ofhow the T box riboswitch antiterminator RNA structurally re-sponds to base pairing with the tRNA acceptor end and of the dis-cernment of the tRNA discriminator base by the antiterminatorelement. The magnesium-dependent results are consistent withthe B. subtilis glyQS 50-UTR in vitro studies in which 5 mM MgCl2

was sufficient to promote the proper folding and binding of thetRNA with the leader sequence, but a minimum of 15 mM MgCl2

was necessary for maximum efficient transcription antitermina-tion [9]. The results of this investigation also support the hypothe-sis that binding of the mh-UCCA with AM1A involves an induced-fit structural rearrangement of the antiterminator model RNA. Thisstructural change accompanies the Mg2+-facilitated base pairing ofthe antiterminator variable base with the tRNA discriminator base.The induced-fit may play a role in the tRNA-induced stabilizationof the antiterminator and the previously-observed resistance ofbound tRNA to displacement.

Acknowledgments

This work was supported by NIH Grant R01-GM61048 and theOffice of the Vice President for Research, Ohio University.

Appendix A. Supplementary data

Supplementary data associated with this article can be found, inthe online version, at doi:10.1016/j.bbrc.2009.09.037.

References

[1] F.J. Grundy, T.M. Henkin, TRNA as a positive regulator of transcriptionantitermination in B. subtilis, Cell 74 (1993) 475–482.

[2] F.J. Grundy, T.M. Henkin, Conservation of a transcription antiterminationmechanism in aminoacyl-tRNA synthetase and amino acid biosynthesis genesin Gram-positive bacteria, J. Mol. Biol. 235 (1994) 798–804.

[3] T.M. Henkin, B.L. Glass, F.J. Grundy, Analysis of the Bacillus subtilis tyrS gene:conservation of a regulatory sequence in multiple transfer-RNA synthetasegenes, J. Bacteriol. 174 (1992) 1299–1306.

J.A. Means et al. / Biochemical and Biophysical Research Communications 389 (2009) 616–621 621

[4] A.G. Vitreschak, A.A. Mironov, V.A. Lyubetsky, M.S. Gelfand, Comparativegenomic analysis of T-box regulatory systems in bacteria, RNA 14 (2008) 717–735.

[5] F.J. Grundy, S.M. Rollins, T.M. Henkin, Interaction between the acceptor end oftRNA and the T-box stimulates antitermination in the Bacillus subtilis tyrS gene:a new role for the discriminator base, J. Bacteriol. 176 (1994) 4518–4526.

[6] S.M. Rollins, F.J. Grundy, T.M. Henkin, Analysis of cis-acting sequence andstructural elements required for antitermination of the Bacillus subtilis tyrSgene, Mol. Microbiol. 25 (1997) 411–421.

[7] F.J. Grundy, S.E. Hodil, S.M. Rollins, T.M. Henkin, Specificity of tRNA–mRNAinteractions in Bacillus subtilis tyrS antitermination, J. Bacteriol. 179 (1997)2587–2594.

[8] F.J. Grundy, W.C. Winkler, T.M. Henkin, TRNA-mediated transcriptionantitermination in vitro: codon-anticodon pairing independent of theribosome, Proc. Natl. Acad. Sci. USA 99 (2002) 11121–11126.

[9] M.R. Yousef, F.J. Grundy, T.M. Henkin, Structural transitions induced by theinteraction between tRNAGly and the Bacillus subtilis glyQS T box leader RNA, J.Mol. Biol. 349 (2005) 273–287.

[10] F.J. Grundy, M.R. Yousef, T.M. Henkin, Monitoring uncharged tRNA duringtranscription of the Bacillus subtilis glyQS gene, J. Mol. Biol. 346 (2005) 73–81.

[11] T.D. Bradrick, J.P. Marino, Ligand-induced changes in 2-aminopurinefluorescence as a probe for small molecule binding to HIV-1 TAR RNA, RNA10 (2004) 1459–1468.

[12] M.E. Gondert, R.A. Tinsley, D. Rueda, N.G. Walter, Catalytic core structure of thetrans-acting HDV ribozyme is subtly influenced by sequence variation outsidethe core, Biochemistry 45 (2006) 7563–7573.

[13] D.A. Harris, D. Rueda, N.G. Walter, Local conformational changes in thecatalytic core of the trans-acting hepatitis delta virus ribozyme accompanycatalysis, Biochemistry 41 (2002) 12051–12061.

[14] D.A. Lafontaine, T.J. Wilson, Z.-Y. Zhao, D.M.J. Lilley, Functional grouprequirements in the probable active site of the VS ribozyme, J. Mol. Biol. 323(2002) 23–34.

[15] K. Lang, R. Rieder, R. Micura, Ligand-induced folding of the thiM TPP riboswitchinvestigated by a structure-based fluorescence spectroscopic approach,Nucleic Acids Res. 35 (2007) 5370–5378.

[16] C. Clerte, K.B. Hall, Global and local dynamics of the U1A polyadenylationinhibition element (PIE) RNA and PIE RNA-U1A complexes, Biochemistry 43(2004) 13404–13415.

[17] M. Kaul, C.M. Barbieri, D.S. Pilch, Fluorescence-based approach for detectingand characterizing antibiotic-induced conformational changes in ribosomalRNA: comparing aminoglycoside binding to prokaryotic and eukaryoticribosomal RNA sequences, J. Am. Chem. Soc. 126 (2004) 3447–3453.

[18] R.K. Neely, D. Daujotyte, S. Grazulis, S.W. Magennis, D.T.F. Dryden, S.Klimasauskas, A.C. Jones, Time-resolved fluorescence of 2-aminopurine as aprobe of base flipping in M.HhaI–DNA complexes, Nucleic Acids Res. 33 (2005)6953–6960.

[19] D. Tleugabulova, L.J. Reha-Krantz, Probing DNA polymerase–DNA interactions:examining the template strand in exonuclease complexes using 2-aminopurine fluorescence and acrylamide quenching, Biochemistry 46(2007) 6559–6569.

[20] E.Y.M. Bonnist, A.C. Jones, Long-wavelength fluorescence from 2-aminopurine-nucleobase dimers in DNA, ChemPhysChem 9 (2008) 1121–1129.

[21] E.L. Rachofsky, R. Osman, J.B.A. Ross, Probing structure and dynamics of DNAwith 2-aminopurine: effects of local environment on fluorescence,Biochemistry 40 (2001) 946–956.

[22] I. Zagórowska, R.W. Adamiak, 2-Aminopurine labeled RNA bulge loops.Synthesis and thermodynamics, Biochimie 78 (1996) 123–130.

[23] S.R. Kirk, N.W. Luedtke, Y. Tor, 2-Aminopurine as a real-time probe ofenzymatic cleavage and inhibition of hammerhead ribozymes, Bioorg. Med.Chem. 9 (2001) 2295–2301.

[24] M.A. O’Neill, J.K. Barton, 2-Aminopurine: a probe of structural dynamics andcharge transfer in DNA and DNA:RNA hybrids, J. Am. Chem. Soc. 124 (2002)13053–13066.

[25] M.J. Rist, J.P. Marino, Association of an RNA kissing complex analyzed using 2-aminopurine fluorescence, Nucleic Acids Res. 29 (2001) 2401–2408.

[26] J.A. Means, S. Wolf, A. Agyeman, J.S. Burton, C.M. Simson, J.V. Hines, T boxriboswitch antiterminator affinity modulated by tRNA structural elements,Chem. Biol. Drug Des. 69 (2007) 139–145.

[27] A.R. Nelson, T.M. Henkin, P.F. Agris, TRNA regulation of gene expression:interactions of an mRNA 50-UTR with a regulatory tRNA, RNA 12 (2006) 1254–1261.

[28] M.S. Gerdeman, T.M. Henkin, J.V. Hines, In vitro structure–function studies ofthe Bacillus subtilis tyrS mRNA antiterminator: evidence for factor-independenttRNA acceptor stem binding specificity, Nucleic Acids Res. 30 (2002) 1065–1072.

[29] K.D. Jack, J.A. Means, J.V. Hines, Characterizing riboswitch function:identification of Mg2+ binding site in T box antiterminator RNA, Biochem.Biophys. Res. Commun. 370 (2008) 306–310.

[30] J.A. Means, S. Katz, A. Nayek, R. Anupam, J.V. Hines, S.C. Bergmeier, Structure–activity studies of oxazolidinone analogs as RNA-binding agents, Bioorg. Med.Chem. Lett. 16 (2006) 3600–3604.

[31] C. Kao, M. Zheng, S. Rudisser, A simple and efficient method to reducenontemplated nucleotide addition at the 30 terminus of RNAs transcribed byT7 RNA polymerase, RNA 5 (1999) 1268–1272.

[32] J.F. Milligan, D.R. Groebe, G.W. Witherell, O.C. Uhlenbeck, Oligoribonucleotidesynthesis using T7 RNA-polymerase and synthetic DNA templates, NucleicAcids Res. 15 (1987) 8783–8798.

[33] J.F. Milligan, O.C. Uhlenbeck, Synthesis of small RNAs using T7 RNA-polymerase, Methods Enzymol. 180 (1989) 51–62.

[34] C.R. Guest, R.A. Hochstrasser, L.C. Sowers, D.P. Millar, Dynamics of mismatchedbase-pairs in DNA, Biochemistry 30 (1991) 3271–3279.

[35] K.H. Scheit, H.R. Rackwitz, Synthesis and physicochemical properties of twoanalogs of poly(dA): poly(2-aminopurine-9-b-D-deoxyribonucleotide) andpoly 2-amino-deoxyadenylic acid, Nucleic Acids Res. 10 (1982) 4059–4069.

[36] D.C. Ward, E. Reich, L. Stryer, Fluorescence studies of nucleotides andpolynucleotides. I. Formycin, 2-aminopurine riboside, 2, 6-diaminopurineriboside, and their derivatives, J. Biol. Chem. 244 (1969) 1228–1237.

[37] R.K. Neely, A.C. Jones, Influence of base dynamics on the conformationalproperties of DNA: observation of static conformational states in rigidduplexes at 77 K, J. Am. Chem. Soc. 128 (2006) 15952–15953.

[38] O.J.G. Somsen, L.B. Keukens, M.N. de Keijzer, A. van Hoek, H. van Amerongen,Structural heterogeneity in DNA: temperature dependence of 2-aminopurinefluorescence in dinucleotides, ChemPhysChem 6 (2005) 1622–1627.

[39] L. Tsujikawa, M.G. Strainic, H. Watrob, M.D. Barkley, P.L. DeHaseth, RNApolymerase alters the mobility of an A-residue crucial to polymerase-inducedmelting of promoter DNA, Biochemistry 41 (2002) 15334–15341.

[40] M.S. Gerdeman, T.M. Henkin, J.V. Hines, Solution structure of the Bacillussubtilis T-box antiterminator RNA: seven nucleotide bulge characterized bystacking and flexibility, J. Mol. Biol. 326 (2003) 189–201.

[41] R.K. Neely, S.W. Magennis, S. Parsons, A.C. Jones, Photophysics and X-raystructure of crystalline 2-aminopurine, ChemPhysChem 8 (2007) 1095–1102.

[42] M. Rist, H.A. Wagenknecht, T. Fiebig, Exciton and excimer formation in DNA atroom temperature, ChemPhysChem 3 (2002) 704–707.

[43] T.J. Su, B.A. Connolly, C. Darlington, R. Mallin, D.T.F. Dryden, Unusual 2-aminopurine fluorescence from a complex of DNA and the EcoKImethyltransferase, Nucleic Acids Res. 32 (2004) 2223–2230.

[44] T. Lenz, E.Y.M. Bonnist, G. Pljevaljcic, R.K. Neely, D.T.F. Dryden, A.J. Scheidig,A.C. Jones, E. Weinhold, 2-Aminopurine flipped into the active site of theadenine-specific DNA methyltransferase M.Taql: crystal structures and time-resolved fluorescence, J. Am. Chem. Soc. 129 (2007) 6240–6248.