Comparison of a PreQ₁ Riboswitch Aptamer in Metabolite-bound and Free States with Implications for Gene Regulation

Abstract

Riboswitches are RNA regulatory elements that govern gene expression by recognition of small molecule ligands via a high affinity aptamer domain. Molecular recognition can lead to active or attenuated gene expression states by controlling accessibility to mRNA signals necessary for transcription or translation. Key areas of inquiry focus on how an aptamer attains specificity for its effector, the extent to which the aptamer folds prior to encountering its ligand, and how ligand binding alters expression signal accessibility. Here we present crystal structures of the preQ₁ riboswitch from Thermoanaerobacter tengcongensis in the preQ₁-bound and free states. Although the mode of preQ₁ recognition is similar to that observed for preQ₀, surface plasmon resonance revealed an apparent K_D of 2.1 ± 0.3 nm for preQ₁ but a value of 35.1 ± 6.1 nm for preQ₀. This difference can be accounted for by interactions between the preQ₁ methylamine and base G5 of the aptamer. To explore conformational states in the absence of metabolite, the free-state aptamer structure was determined. A14 from the ceiling of the ligand pocket shifts into the preQ₁-binding site, resulting in “closed” access to the metabolite while simultaneously increasing exposure of the ribosome-binding site. Solution scattering data suggest that the free-state aptamer is compact, but the “closed” free-state crystal structure is inadequate to describe the solution scattering data. These observations are distinct from transcriptional preQ₁ riboswitches of the same class that exhibit strictly ligand-dependent folding. Implications for gene regulation are discussed.

Introduction

Riboswitches are small, structured non-coding RNA molecules commonly located in the 5′ leader sequences of mRNA. These motifs are usually organized into a high affinity aptamer domain that binds a ligand and a downstream expression platform that possesses sequences necessary to direct transcription or translation (1). It has been estimated that as many as 4.1% of bacterial genes are controlled by such regulatory RNA elements (2, 3), with emerging evidence suggesting that some are susceptible to antimicrobial agents (4, 5).

At present, structures of nearly a dozen distinct classes of riboswitch aptamers have been solved in complex with their cognate ligands (6–8), but only three have been reported in the ligand-free state (8–11). Despite this disparity, comparisons of bound and free-state aptamers have provided valuable insight into the means by which metabolite binding transmits chemical binding information in the aptamer to distal expression platform signals that activate or attenuate transcription or translation (7, 11–13). Toward understanding this problem, we undertook a structural and functional analysis of a preQ₁ riboswitch from Thermoanaerobacter tengcongensis in the metabolite-bound and free states.

PreQ₁ is a metabolic intermediate in the synthetic pathway that produces the hypermodified guanine nucleotide, queuosine (Q)² (Fig. 1A), which is ubiquitous in the eubacterial and eukaryal domains of life (14). Whereas bacteria synthesize preQ₁ de novo in a multistep reaction that starts with GTP (15, 16), eukaryotes are devoid of this pathway and acquire Q from dietary sources or gut flora. A specialized enzyme called tRNA-guanine transglycosylase is then used to insert the Q base at position 34 of select tRNAs (17). The Q modification has been invoked as a source of translational fidelity and has been implicated in bacterial virulence (18) as well as tyrosine bioynthesis in animals (19). The prevalence of Q^(minus)-tRNA is correlated with neoplastic transformation and has been proposed as a metric to grade malignancies (20).

FIGURE 1.

Schematic diagrams of queuosine-precursor metabolites and preQ₁ riboswitch organization. A, relationship between GTP and the hypermodified base queuosine, whose synthesis proceeds through preQ₀ (7-cyano-7-deazaguanine) and preQ₁ (7-aminomethyl-7-deazaguanine) intermediates via the nitrile reductase QueF. In eubacteria, TGTase (tRNA transglycosylase) inserts preQ₁ into the wobble position of specific tRNAs, where it is modified further to give Q (14). B, secondary structure and nucleotide interaction diagram for the T. tengcongensis aptamer adapted from Ref. 22. The color code is as follows. Red, bases involved in Watson-Crick pairs (triple or double solid lines); orange, A-rich region; purple, strictly conserved, non-canonical base interactions with metabolite; slate, decaloop bases engaged in non-canonical pairing and base triples; teal bases are bulged. Q, the metabolite. Interactions are as follows. W, Watson-Crick face; S, sugar edge; H, Hoogsteen edge. The RBS is highlighted in yellow.

Bacterial genes involved in preQ₁ biosynthesis and cellular import were found to be regulated by a small, 34-nucleotide RNA motif known as the class 1 preQ₁ riboswitch (21). At present, this is the smallest known riboswitch aptamer, which was predicted to adopt a stem-loop structure followed by an A-rich tail (21). Subclasses of preQ₁ aptamers were labeled type I and II based on distinct consensus sequences, although their ligand specificities were reported to be the same (21). Subsequently, three independent laboratories described high resolution preQ₁ aptamer structures. The type I aptamer from T. tengcongensis was solved in the presence of preQ₀ (22) and is involved in translational regulation. Its organization is notable because a significant portion of the ribosome-binding site (RBS) is located within the aptamer domain. By contrast, the type II aptamer from Bacillus subtilis was solved in complex with preQ₁ and exhibits spatially distinct aptamer and expression platform domains responsible for transcriptional regulation (23, 24). Nonetheless, both aptamer types are organized as H-type pseudoknots in which ∼50% of bases engage in triples or quartet interactions (Fig. 1B). Solution analysis of type II aptamers from different species revealed folding that was strictly dependent on the presence of ligand with large structural changes observed upon preQ₁ binding (23, 25, 26). Global conformational changes have also been invoked for the thiamine pyrophosphate and glycine riboswitches (27–30).

In this investigation, we conducted crystallographic and solution small angle x-ray scattering (SAXS) analyses of the T. tengcongensis riboswitch in the preQ₁-bound and free states. A parallel SAXS analysis was conducted on the type II Fusobacterium nucleatum preQ₁ riboswitch, which has demonstrated ligand-dependent folding properties (25, 26). We then used surface plasmon resonance to measure the affinity of ligand interactions with the T. tengcongensis aptamer and to identify key affinity determinants as well as the basis for preQ₁ versus preQ₀ selectivity. The results suggest a mechanism of translational attenuation in which metabolite binding is coupled structurally to the solvent accessibility of the RBS. The apparent ability of T. tengcongensis aptamer to adopt a compact, folded conformation in the free state stands in sharp contrast to the type II transcriptional aptamers.

EXPERIMENTAL PROCEDURES

Preparation of RNA Aptamers and Crystallization

All RNA strands were synthesized by Dharmacon Inc. (Fayette, CO). The 33-mer used in crystallization (Fig. 1B) was deprotected, purified by HPLC, and desalted as described (31). The 34-mer aptamer from F. nucleatum harbored a double mutation (26) and was prepared similarly. Strands used for surface plasmon resonance (SPR) were prepared with a 5′-biotin modification and comprised the wild type sequence or point variations in Table 3.

TABLE 3.

Affinity constants of the T. tengcongensis preQ₁ riboswitch and its variants for metabolites

	kon × 104	S.E. × 102	S.D. × 104	koff × 10−4	S.E. × 10−6	S.D. × 10−4	KDa	S.D.	t½	Krel (KD(mut)/KD(WT preQ1)
	m−1s−1			s−1			nm		h
Wild type preQ1	7.77	3.55	3.45	1.53	1.00	0.47	2.05	0.29	1.4	1
Wild type preQ0	0.65	0.27	0.13	2.22	1.04	0.14	35.10	6.07	0.9	17
G11dG preQ1	8.47	2.94	3.71	1.42	0.96	0.58	1.69	0.22	1.6	0.8
G11dG preQ0	0.65	0.20	0.10	1.83	1.07	0.27	28.90	7.7	1.1	14
G5DAP/C16U preQ1	—b	—	—	—	—	—	2.01 × 103	3.32 × 102	—	981
G5DAP/C16U preQ0	—	—	—	—	—	—	1.63 × 103	1.17 × 102	—	795
U6C preQ1	—	—	—	—	—	—	>2.74 × 105	3.05 × 104	—	—
U6C preQ0	—	—	—	—	—	—	>1.99 × 105	2.01 × 104	—	—
A29Pur preQ1	—	—	—	—	—	—	1.84 × 104	3.34 × 103	—	8976
A29Pur preQ0	—	—	—	—	—	—	1.80 × 104	8.42 × 102	—	8781

^a Here we use the nomenclature K_D with the clarification that it is an apparent K_D.

^b —, not determined (apparent K_D obtained from equilibrium analysis) or an estimate that precludes K_rel calculation.

Enzymatic Synthesis of PreQ₀ and PreQ₁

Synthetic preQ₀ was produced as described (15, 32). PreQ₀ was converted to preQ₁ by the E. coli enzyme QueF, which was purified as described (33). The reaction for the enzymatic conversion contained 0.050 m potassium phosphate (pH 7.4), 0.10 m KCl, 0.5 mm NADPH, 0.5 mm preQ₀, and 10 μm QueF in a final volume of 0.40 liter. The mixture was stirred gently for 8 h at 37 °C and filtered through a YM-10 (Amicon) membrane under N₂(g) to remove QueF. The filtrate was lyophilized, resuspended in 0.20 liters of methanol, and stirred for 1 h to dissolve preQ₁. Undissolved material was removed by filtration, and methanol was removed by rotary evaporation until only a yellow solid remained. The solid was dissolved in 10 ml of water (pH 3.8) and loaded onto a 60-ml CM Sepharose Fast Flow column (2.6 × 11.3 cm; GE Healthcare) in water (pH 3.8). The column was rinsed with 0.10 liter of water (pH 3.8), and preQ₁ was eluted with a linear gradient extending to 0.5 m HCl in a volume of 0.50 liter. Fractions (5 ml) were collected and analyzed for the presence of preQ₁ by LC-MS using the following protocol. 25 μl of material was injected onto a 4.6 × 250-mm Eclipse XDB-C18 column (Agilent) that had been pre-equilibrated in water. PreQ₁ was eluted by developing the column with a gradient to 40% acetonitrile over 40 min at a flow rate of 0.3 ml/min. The elution was monitored by UV-visible and MS detection. UV-visible spectra were obtained from 220 to 500 nm using a ThermoFinnigan Surveyor photodiode array detector. Mass spectra were obtained in positive mode, scanning the m/z range of 100–300 atomic mass units using an electrospray ionization-equipped LCQ ThermoFinnigan Deca XP mass spectrometer. The instrument was set at a 6 V ionization energy and a 200 °C ion source temperature. Fractions containing preQ₁ were pooled, lyophilized, and dissolved in 1 ml of water. The concentration of preQ₁ was determined by UV absorbance using extinction coefficients published for 7-deazaguanine at pH 6.8 (34).

Crystallization and X-ray Structure Determination

The free-state riboswitch was crystallized by suspending lyophilized RNA in 0.010 m sodium cacodylate buffer, pH 7.0, to a concentration of 0.5 mm. A total of 20 μl was heated to 65 °C for 2 min before cooling to 30 °C in an aluminum block, followed by the addition of MgCl₂ to a final concentration of 0.010 m. The sample was heated again to 65 °C and held for 3 min, followed by slow cooling in a 0.5-liter water bath to 20 °C over 2 h. Crystallization trials were set up using hanging drop vapor diffusion in which 2 μl of RNA was added to 2 μl of mother liquor comprising 2.0 m Li₂SO₄, 0.050 m MES, pH 6.0, 0.003 m MgSO₄ and 0.002 m spermine at 20 °C. Crystals grew as hexagonal rods to 0.4 × 0.15 × 0.15 mm in 6–8 weeks. Metabolite-bound crystals were prepared as described (22) using preQ₁ in place of preQ₀. Crystals were cryoprotected in equal mixtures of Paratone-N and silicone oils (Hampton Research) and flash frozen (22).

X-ray diffraction data were collected on site at the Cornell High Energy Synchrotron Source (CHESS) (Ithaca, NY), and remotely at the Stanford Synchrotron-Radiation Lightsource (SSRL) (Menlo Park, CA) using the Blu-Ice and Web-Ice interfaces (35). Intensity data were reduced using HKL2000 (36). Structures were solved by difference Fourier methods in PHENIX (37) using the preQ₀ structure (Protein Data Bank entry 3gca) as a starting model. Both structures were subjected to manual rebuilding in COOT (38). The free-state structure was subjected to Cartesian simulated annealing with rebuilding into composite omit maps. Intensity and refinement statistics are provided in Table 1. Figure graphics were generated in PyMOL (39). Solvent accessibility was calculated with AREAIMOL (40).

TABLE 1.

X-ray data reduction and refinement statistics

	PreQ1-bound + Mg2	Free state (Mg2+ only)
X-ray data reduction statistics
Protein Data Bank accession code	3Q50	3Q51
Beamline	CHESS A-1	SSRL 9–2
Wavelength (Å)	0.98	0.98
Space group	P6322	P6322
Unit cell (Å)	a = 110.8, c = 59.6	a = 116.6, c = 57.9
Total observations	38,982	41,267
Unique observations	6041	5771
Resolution (Å)	22-2.75 (2.80-2.75)a	27.8-2.85 (2.95-2.85)
Completeness (%)	99 (96)	99.2 (100)
〈Redundancy〉	6.5 (5.1)	7.2 (8.4)
I/σ(I)	24.7 (2.1)	20.6 (6.7)
Rsym (%)b	7.1 (43.6)	7.5 (35.7)
Refinement statistics
Rcryst/Rwork/Rfree (%)c,d,e	21.4/21.0/25.2	23.1/22.6/27.2
Geometry
Root mean square bonds (Å)	0.007	0.006
Root mean square angles (degrees)	1.689	0.844
Coordinate error (Å)	0.35	0.40
No. of atoms/B-factors
RNA	702/56.6 Å2	680/56.7
Metabolite	13/47.7 Å2	None
SO42−	20/67.5 Å2	5/51.8 Å2
Mg2+	None	1/60.3 Å2
Anisotropy correction	B11:12.6 B33:−25.2	B11:10.6 B33:−21.2

^a Values in parentheses represent high resolution statistics.

^b R_sym = [Σ|I(h)_j − 〈I(h)〉|/Σ|I(h)_j|] × 100, where I(h)_j is the intensity of the jth reflection of type h, and 〈I(h)〉 is the average intensity of a reflection of type h.

^c R_cryst represents a final cycle of refinement without cross-validation using the residual target and all reflections.

^d R_work = [Σ_hkl|(F_o) − k(F_c)|]/[Σ_hkl|(F_o)|] × 100, where k is a scale factor and the test set has been removed.

^e R_free is defined as the R_work calculated for 9% of the x-ray data selected randomly and excluded from refinement.

SAXS

Experiments were conducted at the G1 station of CHESS. Using a wavelength of 1.29 Å, scattered X-rays were recorded on a CCD detector at a sample-to-detector distance of 94.2 cm. This configuration gave an accessible q range from 0.025 to 0.35 Å⁻¹, where q = 4πsinθ λ⁻¹, in which 2θ is the scattering angle and λ is the radiation wavelength. Samples of the preQ₁ aptamer were prepared in the presence or absence of metabolite in 0.010 m sodium cacodylate, pH 6.0, with 0.010 m MgCl₂. Three sample concentrations were utilized, including 0.075, 0.20, and 0.55 mm. Microcentrifuged samples were transferred to disposable cells (41) held at 20 °C. Buffer blanks were collected for each cell. Exposures were recorded as 3 × 3 s followed by 3 × 9 s series. Scattering data for the 3 × 9 s exposures were integrated by Data Squeeze, version 2.20 (P. Heiney) to yield the respective one-dimensional intensity profiles. No evidence of radiation damage was observed based on a comparison of initial versus final exposures. Buffer corrections and extrapolation to zero concentration were accomplished using PRIMUS (43); it has been noted that extrapolation to infinite dilution can circumvent problems with interparticle interference that can cause inaccurate measurements (44). Kratky plots were generated for the full range of data, and Guinier plots were calculated with PRIMUS. GNOM was used to produce the paired distance distribution functions, which were optimized to meet visual perception criteria as described (45). Dimensions of crystal structures and fits between experimental and crystallographic scattering profiles were calculated using CRYSOL (46); additional dimensions were obtained by use of MOLEMAN (47). Relevant SAXS parameters are reported in Table 2.

TABLE 2.

Solution dimensions from small angle x-ray scattering

	RgG	RgP	rmax
	Å	Å	Å
Wild type preQ1 + Mg2+	16.6 ± 0.02	16.5 ± 0.01	51 ± 5
Wild type preQ0 + Mg2+	16.9 ± 0.02	16.9 ± 0.01	57 ± 6
Wild type Mg2+	17.0 ± 0.02	17.0 ± 0.01	57 ± 6
F. nucleatum preQ1 + Mg2+	19.3 ± 0.03	19.3 ± 0.01	64 ± 6
F. nucleatum Mg2+	31.0 ± 0.07	30.9 ± 0.02	107 ± 11
Crystal coordinates, preQ0/Q1		16.9a	50b
Crystal coordinates, free state		17.0a	52b

^a Calculation considers the solvent shell from CRYSOL (46).

^b From MOLEMAN (47), including 1.6 Å for a monolayer shell of water on the molecular surface (65).

SPR

NeutrAvidin-conjugated CM5 chips were prepared on a Biacore T100 instrument (Biacore Inc., Piscataway, NJ) using reagents from the manufacturer. Biotinylated free-state preQ₁ aptamers and variants thereof (Table 3) were suspended in running buffer (0.010 m sodium cacodylate, pH 7.0, and 0.003 m MgCl₂) to a concentration of 400 nm. RNA aptamers were injected over NeutrAvidin-conjugated flow cells to give a surface density of 3800–4000 response units, with the reference flow cell having only NeutrAvidin. For kinetic experiments, the association phase consisted of 180-s injections of preQ₁ or preQ₀ conducted at concentrations ranging from 1 to 250 nm at 75 μl/min; dissociation was monitored for 600 s. Regeneration was conducted using 3 m guanidine HCl for 180 s followed by a 120-s incubation in running buffer flowing at 30 μl/min. Equilibrium binding experiments were carried out as follows. Injections of metabolite lasted for 240 s at 30 μl/min and were conducted with concentrations ranging from 0.470 to 400 μm. A dissociation phase of 400 s ensued in running buffer. Regeneration was accomplished using 3 m guanidine HCl for 90 s, followed by a 160-s stabilization period in running buffer at 30 μl/min. All measurements were made in duplicate at 25 °C with a data collection rate of 10 Hz. Experimental data were processed using Biacore T100 Evaluation software version 2.2 (GE Healthcare) and the double referencing method (48) to remove instrumental and bulk shift effects. The buffer-subtracted sensorgrams for the kinetic binding data were then fit to a 1:1 binding interaction model to determine the rate constants (k_on and k_off) and the apparent equilibrium binding constant (K_D). To evaluate the steady-state equilibrium binding data, the response during the equilibrium binding phase (R_eq) was calculated by averaging the response over 5 s just before the end of the metabolite injection. Data were fit using non-linear curve fitting using PRISM4 (GraphPad, Inc.) to a one-site binding equation to determine K_D.

RESULTS

Quality Indicators and Overall Fold of the PreQ₁-bound Riboswitch Aptamer

To understand the basis of metabolite specificity and gene regulation by the class 1, type I T. tengcongensis preQ₁ translational riboswitch, we undertook a structure determination in the presence of the primary target ligand, preQ₁ (Fig. 1A). The entire 33-mer RNA aptamer was observed in electron density maps as well as the preQ₁ ligand, whose omit electron density map is representative of the overall quality of the model (Table 1 and Fig. 2A). The hydrogen bonding pattern, global fold, and location of the metabolite closely matched features reported previously for the preQ₀-bound state (Figs. 1B and 2B) (22). The quality of the 2.75 Å resolution preQ₁-bound structure is indicated by its R_work and R_free values of 21.0 and 25.2%, respectively (Table 1), which are substantially lower than the preQ₀-aptamer complex, whose R_work and R_free values were 24.5 and 27.2% (22). Despite identical diffraction resolutions, B-factors for the preQ₁-bound state were 57 Ų for RNA atoms and 48 Ų for preQ₁ (Table 1) as compared with B-factors of 76 Ų for RNA atoms and 58 Ų for preQ₀ of the preQ₀-bound state.

FIGURE 2.

Schematic representations of the preQ₁ riboswitch in the free and preQ₁-bound states. A, aptamer interactions with preQ₁ looking down the helical axis of the riboswitch. The “floor” of the binding pocket is shown as a space-filling model; putative hydrogen bonds are shown as dashed lines. The location of the ligand is indicated by an averaged “kicked” (66) or AK (mF_o − DF_c) omit electron density map with σA weighting contoured at 4σ; the ligand (cyan) was removed from the phase calculation. Inset, close-up view of the preQ₁ methylamine in relation to the G11 2′-OH group and the G5 O6 keto moiety. Neighboring groups of G5 are highlighted with a semitransparent circle and labeled; the electron density has been removed for clarity. B, aptamer interactions with preQ₀ based on Protein Data Bank entry 3cga as described previously (22). C, ribbon diagram of the free-state riboswitch. The phosphate backbone is depicted as a teal ribbon with bases colored as in Fig. 1B; inset, a view rotated 90° to accentuate the location of A14 within the ligand-binding pocket. D, schematic diagram depicting A14 of the free-state structure; the purine base was removed from the phase calculation to generate an AK (mF_o − DF_c) omit map as in A.

Quality Indicators and Overall Fold for the Free-state Aptamer

To understand the conformational changes involved in RNA-mediated gene regulation by sequestration of the RBS, we determined the ligand-free T. tengcongensis aptamer structure (Fig. 2C). Compared with the preQ₁-bound state, free-state crystals showed a 7.6% increase in the unit cell volume (Table 1), suggestive of significant structural differences. Although the overall fold of the free-state structure is comparable with the compact 48 Å × 28 Å × 15 Å preQ₁-bound state, changes occurred in the RNA backbone flanking the metabolite-binding pocket. No electron density was observed for A13 of the decaloop, indicated as a dashed line in the backbone of Fig. 2C, and the quality of the electron density in this region was generally poorer than that of either ligand-bound state. The overall RNA B-factors were comparable at 57 Ų, but the R_work and R_free values were slightly worse than the preQ₁-bound state (Table 1). Significantly, the electron density was of sufficient quality and resolution to observe conformational changes in the decaloop at C15 and A14 that are consistent with changes resulting from ligand-free crystallization as opposed to crystal packing because this area is devoid of lattice contacts. Reduced bias omit-electron density maps, such as that calculated for the A14 base of the free-state structure (Fig. 2D), indicate the quality of the model in this region. The level of change relative to the coordinate error (Table 1) is sufficiently large to permit a meaningful structural comparison on both global and local levels.

Comparison of the PreQ₁-bound and Free-state Crystal Structures Reveals a Network of Localized Conformational Changes

To contrast the preQ₁-bound and free state aptamers, an all-atom superposition was conducted (Fig. 3A). Although only a few nucleotides differed significantly in their relative positions, this was sufficient to elevate the root mean square displacement of atoms to 1.7 Å. The maximum displacement was 7.4 Å, which corresponded to the N1 imino of A14. The nucleotide at this position rotates out of a stable G11 to A14 (N3-amino amino-N1) pair flanking the ligand pocket to occupy the preQ₁ binding site (Fig. 3B, compare dark blue A14 to light blue A14). Other pronounced changes cluster in the pseudoknotted decaloop surrounding the preQ₁ binding site. C15 rotates away from the binding pocket, a 6.6-Å movement of N3, from a location where it base-pairs with preQ₁ (Fig. 2, A versus D, and Fig. 3, dark blue C15 versus red). By contrast, U12 shifts by 4.4 Å toward the binding pocket closer to the position vacated by A14. Finally, base A32 moves away from the binding pocket in the free-state structure by 2.3 Å. These changes appear interconnected to the A14 shift, which has the effect of eliminating the continuous base stack that links A32 and G33 of the RBS to preQ₁ in the aptamer core in the ligand-bound state.

FIGURE 3.

All-atom superposition of the respective free-state and ligand-bound structures. A, schematic stereo view of the global, free-state structure (colored as in Fig. 2) superimposed upon the preQ₁-bound structure (dark blue) using LSQKAB (42). Black arrows indicate the movements of bases in the preQ₁-bound structure relative to those of the free state. The start of the ribosome-binding site at A32 is labeled RBS. The RBS portion of the aptamer is highlighted in yellow. The preQ₁ metabolite is labeled Q₁. A ribbon passing through the phosphate backbone is shown in teal (free state) and blue (preQ₁-bound). B, close-up stereo diagram of A (rotated +90° along the x axis) depicting the superposition of select decaloop nucleotides. Putative hydrogen bonds to preQ₁ in the ligand-bound structure are drawn as broken lines. Nucleotides that superimpose with a root mean square deviation of >2.0 Å are labeled twice for the respective ligand-bound (blue) or free state (colored as in Fig. 2C) structures. Base G11 was rendered with thin bonds for clarity; here only the preQ₁-bound state is shown because of its close similarity to the free-state structure.

Molecular Recognition of PreQ₁ and Closure of the Binding Pocket in the Free State

Although the mode of preQ₀ binding has been described for the T. tengcongensis riboswitch aptamer (22), the mode of preQ₁ binding has remained uncertain until now. The compact architecture of the aptamer buries 330 of 337 Ų of the preQ₁ metabolite fortifying the core structure by generating a network of continuously stacked bases that join the RBS at the “top” of the aptamer to the P1 stem (Figs. 1B and 3A). Like preQ₀, the Watson-Crick face of preQ₁ is “read out” by C15 (Fig. 2, A and B), and conserved bases U6 and A29 recognize the minor groove edge of both metabolites. The modified guanine base of the metabolite stacks between the G5-C16 base pair of the P1 stem (Fig. 2A) and bases G11 and A14 (Fig. 3). These interactions have been referred to as the respective “floor” and “ceiling” of the binding pocket (22). However, in the absence of metabolite, the pocket adopts a “closed” conformation whereby binding to preQ₁ is blocked by A14, which spatially overlaps the ligand-binding site (Fig. 3B) and pairs with key bases involved in ligand recognition, most notably U6 and A29 (Fig. 2D). Because A14 forms part of the binding pocket ceiling, its shift in the free state effectively collapses the ligand recognition site and directly increases the solvent accessibility of the RBS from 468 to 511 Ų, which starts at position A32 of the aptamer (Figs. 1B and 3A and supplemental Fig. S1).

Small Angle X-ray Scattering Analysis of the T. tengcongensis and F. nucleatum PreQ₁ Aptamers

An important question regarding the observed preQ₁-bound and free state crystal structures is whether they are consistent with (i) the compactness and (ii) the conformations present in the solution state. To address these questions, we subjected the T. tengcongensis riboswitch to SAXS analysis. We also examined the F. nucleatum 34-mer class 1, type II preQ₁ aptamer as a control, which was chosen for its pronounced ligand-dependent folding (25, 26).

The results revealed smooth intensity distributions with gently changing slopes for both aptamer types (Fig. 4A). Sudden upturns or downturns were not observed, consistent with an absence of aggregation and interparticle repulsion, respectively (49). Visual comparisons indicated that the free-state F. nucleatum aptamer has a more negative slope at low q than the matched sample prepared in the presence of preQ₁ (Fig. 4A, curve 1 versus curve 2). In contrast, the F. nucleatum sample in the presence of preQ₁ is more similar to the T. tengcongensis samples (Fig. 4A, curves 2–4). Based on visual criteria (49), the quality of the scattering profiles was deemed suitable for dimensional analysis.

FIGURE 4.

Small angle x-ray scattering analysis of preQ₁ riboswitches from T. tengcongensis and F. nucleatum in metabolite-bound and free states. A, scattering profiles (thick lines) derived from experimental measurements and those calculated from T. tengcongensis crystal structures (thin gray lines) in Table 1. The samples are free-state F. nucleatum with 10 mm Mg²⁺ (1), preQ₁-bound F. nucleatum with 10 mm Mg²⁺ (2), preQ₁-bound T. tengcongensis with 10 mm Mg²⁺ (3), and free-state T. tengcongensis with 10 mm Mg²⁺ (4). The agreement between the scattering profiles from the experimental SAXS data and crystallographic coordinates of the respective T. tengcongensis preQ₁-bound and free-state crystal structures is given by χ² as defined by CRYSOL (46). B, comparison of the paired distance distribution plots, P(r), for the samples in A. The real-space R_g^P and maximum intraparticle distance, r_max, from these functions are reported in Table 2. The functions were scaled relative to a reference according to the integrated areas of each respective curve.

Comparison of Ligand-induced Compaction of PreQ₁ Aptamers

To evaluate the degree to which the T. tengcongensis translational riboswitch undergoes ligand-dependent compaction, we used three criteria: (i) reduction of radius of gyration (R_g) values derived from both Guinier plots (R_g^G; supplemental Fig. S2B) and paired distance distribution functions (R_g^P; i.e. P(r) plots in Fig. 4B and supplemental Fig. S2C); (ii) reduction of the longest interatomic distance r_max in a sample from P(r) functions; and (iii) the shapes of Kratky plots, which show distinct dependences of momentum transfer-weighted intensity versus q for folded versus unfolded polymers (50) (supplemental Fig. S2D).

Significant differences in ligand-dependent compaction were observed between the F. nucleatum and T. tengcongensis riboswitch aptamers. The F. nucleatum “control” aptamer in the absence of ligand had a significantly larger R_g of 31.0 Å and longer r_max of 107 ± 11 Å compared with its preQ₁-bound state, in which the R_g was 19.3 Å and r_max was 64 ± 6 Å (Fig. 4B and Table 2). The broad, heavy tailed Kratky plots of the free-state F. nucleatum aptamer were consistent with partial folding in the absence of preQ₁ (supplemental Fig. S2D), whereas the bell-shaped, light-tailed profile of the preQ₁-bound aptamer was characteristic of a well folded polymer (supplemental Fig. S2D) (49, 50). Conversely, the T. tengcongensis aptamer appeared as compact in the free state with R_g^G = 17.0 Å and r_max = 57 ± 6 Å, as it is in the preQ₁- and preQ₀-bound states in which the respective R_g^G and r_max values were 51 ± 5 and 16.6 Å and 57 ± 6 and 16.9 Å (Table 2). Kratky profiles for each of the T. tengcongensis samples were comparable with the preQ₁-bound F. nucleatum aptamer, indicative of folded RNA (supplemental Fig. S2D).

Comparison of Crystal Structures with Solution SAXS Data

We then asked whether our crystal structures were representative of solution dimensions and conformational states. The experimentally measured R_g^P and r_max values were in reasonable agreement with those calculated from the respective preQ₁-bound, preQ₀-bound, and free-state crystal structures but with notable differences. The best agreement corresponded to the preQ₁-bound state in which the crystal structure R_g^P and r_max values were 16.9 and 50 Å versus the solution R_g^P and r_max values of 16.5 and 51.5 Å (Table 2). The solution R_g^P values for the preQ₀-bound and free-state samples agreed exactly with crystal structures at 16.9 and 17.0 Å, respectively (Table 2). However, the corresponding solution r_max values for the preQ₀-bound and free-state samples were each 57 Å, which is longer than the corresponding crystal structures, whose values were 50 and 52 Å, respectively. This finding may be the result of greater mobility in solution or conformational restraints imposed on the coordinates by the crystal lattice, which can only accommodate an r_max of ∼52 Å (discussed below).

To assess the agreement between the intensity profiles calculated from crystal structures in Table 1 versus those corresponding to the experimental data, we calculated χ² values (46). The results revealed that the preQ₁-bound crystal structure has a χ² value of 1.7, indicating reasonable agreement between the atom model and the solution conformation (11). In contrast, the free-state crystal structure yielded a χ² of 5.8. Deviations in the intensity fit can be seen at several distinct points along the curve, especially a medium and high q (Fig. 4A, curve 4). This level of discrepancy is strong evidence that the crystallographic model is insufficient to describe the solution conformational ensemble and that differences between the bound and free states are more complex than suggested by a comparison of the corresponding crystal structures, despite similarities in compactness demonstrated by R_g^P and r_max.

Kinetics and Affinity Constants for PreQ₁ and PreQ₀ Binding to the T. tengcongensis Aptamer

PreQ₀ and preQ₁ differ by substitution of a cyano group for a methylamine (Fig. 1A). To evaluate the kinetics and thermodynamics of ligand binding, we measured the rates of binding and dissociation to the T. tengcongensis aptamer using surface plasmon resonance (Fig. 5, A and B). The results revealed that preQ₁ binds with an on-rate of 7.77 × 10⁴ m⁻¹ s⁻¹, which is 12-fold faster than the on-rate for preQ₀ (Table 3). Both metabolites exhibited similar off-rates at 1.53 × 10⁻⁴ s⁻¹ and 2.22 × 10⁻⁴ s⁻¹, respectively. The k_on for the T. tengcongensis aptamer is 8-fold slower than preQ₁ binding to the F. nucleatum 34-mer aptamer, but k_off for the F. nucleatum aptamer is >10³-fold faster (25). The preQ₁ on-rate of the T. tengcongensis aptamer falls into a range of values reported for other translational riboswitches, including the Escherichia coli thiamine pyrophosphate aptamer at 8.66 × 10⁴ m⁻¹ s⁻¹ (51) and the Vibrio vulnificus adenine aptamer at 3.75 × 10⁴ m⁻¹ s⁻¹ (52). By contrast, the off-rate of the T. tengcongensis aptamer is 2 orders of magnitude slower than the TTP (0.043 s⁻¹) and adenine (0.017 s⁻¹) aptamers. At present, the slowness of the T. tengcongensis off-rate is surpassed only by the Vibrio cholerae c-di-GMP riboswitch, a transcriptional regulator whose off-rate is 850-fold slower (53).

FIGURE 5.

Kinetic and affinity analysis of the T. tengcongensis riboswitch aptamer for preQ₁ and preQ₀. A, representative surface plasmon resonance sensorgram for preQ₁ interactions with the wild type aptamer. B, representative surface plasmon resonance sensorgram for preQ₀ interactions with the wild type aptamer. The thick black lines represent the buffer-subtracted data, and gray lines are the global fit obtained from a 1:1 binding model based on the experimental data. The resulting k_on and k_off rates and the apparent equilibrium K_D values are reported in Table 3. C, A29Pur mutant response at equilibrium (R_eq) plotted against the concentration of preQ₁ or preQ₀. D, the U6C mutant as described in C. E, the G5DAP/C16U double mutant as described in C. The plots of R_eq correspond to preQ₁ data points (filled squares) or preQ₀ data points (open squares); error bars are shown. The equilibrium binding constant K_D was determined for each metabolite by curve fitting (black or gray lines) to the respective metabolite R_eq data points as described under “Experimental Procedures.” The apparent K_D values are reported in Table 3.

The T. tengcongensis aptamer apparent equilibrium dissociation constant (K_D) was 2.05 ± 0.29 nm for preQ₁ and 35.10 ± 6.07 nm for preQ₀. The magnitude and affinity trends mirror values reported previously for these metabolites during an analysis of a longer B. subtilis aptamer (20 and 100 nm, respectively) using equilibrium dialysis (21). By contrast, the F. nucleatum aptamer showed a K_D of 283 nm for preQ₁, making the T. tengcongensis aptamer a significantly tighter preQ₁ binder than either of the transcriptional riboswitches in its class.

Using Mutagenesis and SPR to Probe T. tengcongensis Riboswitch Aptamer Binding to Metabolites

Specific predictions about the molecular interactions between the T. tengcongensis aptamer and preQ₁ or preQ₀ can be proposed based on new and prior structural observations (Fig. 2, A and B) (22). To test the importance of key aptamer functional groups consistent with hydrogen bonding to preQ₁ or preQ₀ and to differentiate the modest difference in ligand-binding affinity, we undertook an SPR analysis of specific site-directed mutants that were placed into two categories: (i) those that recognize guanine-like features, and (ii) those that promote preQ₁ binding but not necessarily preQ₀.

Mutagenesis of T. tengcongensis Aptamer Bases That Recognize Guanine-like Features Shared by PreQ₁ and PreQ₀

The first class of mutants examined were those predicted to interact with the guanine-like features held in common by preQ₁ and preQ₀. Prior analysis indicated that position C15 (T. tengcongensis numbering) is a universally conserved base in the class 1 preQ₁ aptamer family and that modifications to this base ablated preQ₁ binding or could be used to alter ligand specificity in a manner consistent with Watson-Crick pairing (21). This finding was corroborated subsequently by three independent structure determinations (22–24), which also revealed the locations of other key groups involved in metabolite recognition. Of special importance are universally conserved bases U6 and A29 because they each recognize the same minor groove edge features of preQ₁ and preQ₀ (Fig. 2, A and B) (22).

Our mutagenesis results revealed that the A29Pur and U6C variants each have detrimental effects on preQ₁ and preQ₀ binding but to varying extents (Fig. 5, C and D). The A29Pur variant is devoid of an exocyclic amine and was predicted to eliminate a single hydrogen bond to the metabolites (Fig. 2, A and B). The resulting K_D values were 18.4 μm for preQ₁ and 18.0 μm for preQ₀, which is nearly 9000-fold worse than wild type binding to preQ₁ (Table 3). Loss of the A29 amine also ablates a core folding interaction with the O₂ keto of C7 (not shown) that may account for a loss in binding affinity for both ligands that is significantly greater than removal of a single hydrogen bond.

The U6C variant was also predicted to eliminate a single hydrogen bond between the aptamer O4 keto group and the N9 imine of the ligand pyrrolo ring (Fig. 2, A and B). However, the U6C change introduces a steric block from replacement of a keto with an amine. This substitution may account for the greater relative loss in binding affinity of this variant compared with A29Pur in which a functional group was eliminated. As expected, the U6C change proved detrimental for preQ₁ and preQ₀, whose K_D values were estimated to be >274 and >199 μm, respectively (Table 3). Like A29Pur, the U6C mutation also has the potential to eliminate two core hydrogen bonds between the Watson-Crick face of U6 and the Hoogsteen edge of A28 (Fig. 2, A and B).

The 2′-OH group of G11 was observed to be within hydrogen bonding distance of the O6 keto group of preQ₀ (Fig. 2B) and was perceived to be a potential contributor to metabolite recognition (22). However, this interaction was not present in the preQ₁-bound state due to an apparent change in the ribose pucker (Fig. 2, A (inset) versus B). Instead, the G11 ribose hydroxyl is oriented close to the methylamine of preQ₁, although it is not in an optimal hydrogen bonding orientation. Mutation of the 2′-OH group of G11 to 2′-deoxy G11 revealed no significant deviation in either preQ₀ or preQ₁ binding but showed slightly more favorable binding properties (Table 3). The fact that the 2′-deoxy G11 variant had no substantial effects on the binding affinity or kinetics of either metabolite appears to rule out a role for this group in metabolite selectivity.

Mutagenesis of T. tengcongensis Aptamer Base Interactions That Promote PreQ₁ Binding but Not PreQ₀

A second category of mutants was designed to identify aptamer functional groups that contribute to the higher affinity of preQ₁ over preQ₀ (2.05 nm versus 35.10 nm, respectively; Table 3). The crystal structure of the T. tengcongensis aptamer bound to preQ₁ provides some insight into this question because the methylamine group is within 4.0 Å of the O6 and N7 groups of conserved base G5 (Figs. 2A (inset) and 3B). However, a sulfate ion bound to the preQ₁ methylamine precludes a cogent interpretation (Fig. 3B).

Previously, the G5 base was shown to be universally conserved in preQ₁ class I aptamers (21). Changes in the J1 stem at position C16 (Fig. 1A) as well as flanking base pairs showed poor responses to ligand binding or failed to elicit metabolite-dependent gene regulation (21). Although no functional analysis was reported for G5, T. tengcongensis crystal structures bound to metabolites suggested that the substitution of 2,6-diaminopurine (DAP) in the context of the double mutation, G5DAP/C16U, would replace the G5 O6 keto group with an N6 amine. A comparable mutational analysis (21) demonstrated that this variant adopts a Watson-Crick-like base pair, which provided confidence that the floor of the binding pocket would remain intact. However, the N6 exocyclic amine from DAP is predicted to disrupt hydrogen bonding to the preQ₁ methylamine, whereas this interaction cannot occur with preQ₀ due to the presence of its linear cyano moiety (Figs. 1A and 2, A and B).

The results revealed K_D values of 2.01 ± 0.33 and 1.63 ± 0.12 μm for preQ₁ and preQ₀, respectively (Fig. 5E and Table 3). The relative losses in affinity suggest a nearly 10³-fold reduction in preQ₁ binding but only a 46-fold loss by preQ₀, thus providing evidence for involvement of the G5 O6 keto in binding preQ₁ but not preQ₀.

DISCUSSION

Riboswitches regulate gene expression by altering accessibility to sequences that signal transcription or translation. Prior structural investigations of the preQ₁ riboswitch focused on type II aptamers that function transcriptionally by altering the interconversion of terminator and anti-terminator stem-loops (23–25). Here we examined the structural and functional properties of a type I aptamer from T. tengcongensis involved in translational regulation. A32 of the aptamer sequence (Fig. 1B) appears to be the start of the RBS based on the Shine-Dalgarno consensus sequence and base-pairing energy with the 16 S rRNA (supplemental Fig. S1). This assignment places the last nucleotide of our construct, G33, as the second base of the RBS (Fig. 1B) and has implications for how the mode of preQ₁ binding by the T. tengcongensis riboswitch can regulate translation.

Crystallographic Conformational Changes Reveal Differences in RBS Accessibility

Comparisons of the preQ₁-bound and ligand-free structures of the T. tengcongensis riboswitch provide the basis to identify ligand-dependent conformational changes that may lead to RNA-mediated gene regulation. A superposition of these structures revealed changes in and around the binding pocket that suggested the absence of metabolite causes a shift of base A14 such that it interacts with preQ₁ binding determinants U6 and A29 (Figs. 2 (A versus D) and 3). The position of A14 in the free state coincides with unstacking of bases A32 and G33 of the RBS from the core fold, which increases their solvent accessibility relative to the preQ₁-bound state. Although this difference is modest in the context of the crystal lattice, the tangible increase in accessibility (Fig. 3A) is illustrative of how the 16 S rRNA of the ribosome could gain greater access to the RBS in the free-state structure, thereby resulting in a “gene-on” state.

Modes of PreQ₁ and PreQ₀ Binding

The mode of metabolite binding by members of the class I preQ₁ riboswitch family appears to be conserved insofar as the B. subtilis (type II) and T. tengcongensis (type I) aptamer structures utilize a common set of core bases for recognition. Our functional analysis supports the importance of U6 and A29 in recognition of both the preQ₀ and preQ₁ metabolites (Table 3), thus corroborating crystallographic observations. Efforts to understand the molecular basis for the affinity preference of preQ₁ over preQ₀ led us to examine the G5DAP/C16U double mutant. Our crystal structure shows close proximity between the preQ₁ methylamine and the O6 keto of G5 (Fig. 2A, inset). However, a sulfate ion from the mother liquor is bound to the methylamine group of preQ₁ (Fig. 3B), thereby obfuscating its potential for recognition by the aptamer. We hypothesized that substitution of the G5 O6 keto with a DAP N6 amine would sterically occlude binding of the preQ₁ methylamine and prevent formation of a hydrogen bond. Differences in the K_D values between preQ₁ and preQ₀ correspond to a free energy change of 1.68 kcal/mol. This modest difference suggests no more than 1–2 neutral hydrogen bonds (54). Our results for this mutant suggested a slight favorability in preQ₀ binding over preQ₁, which seems considerable given the 17-fold greater affinity that wild type exhibits for preQ₁ over preQ₀ (Table 3). Such a trend supports the expectation that the G5DAP mutation is more detrimental for preQ₁ than preQ₀ due to the steric block created by the DAP N6 amine. Although the deletion of the G5 keto might appear to be a more conservative probe for hydrogen bonding, we examined a G52AP (2-amino-purine) mutant devoid of an O6 keto moiety and measured the K_D for preQ₁ at >914 μm (data not shown). Changes in mutant affinity for metabolites relative to wild type suggest that substitutions in the aptamer core have additional long range effects that go beyond energy differences expected for loss of 1–2 core hydrogen bonds.

A comparative structural analysis provided further insight into the basis of preQ₁ versus preQ₀ specificity. Commonalties are apparent in the mode of preQ₁ recognition by conserved bases shared by the T. tengcongensis aptamer and the known B. subtilis aptamer structures. This makes it possible to understand the basis for preQ₁ specificity for all class 1 preQ₁ riboswitch family members. In each independent B. subtilis structure, the methylamine of preQ₁ is sufficiently close to the O6 keto and N7 imino of G5 that the metabolite is within hydrogen bonding distance of these groups (23, 24). These interactions are not present when preQ₀ is bound to the T. tengcongensis aptamer (22) and would account for the observed energy difference that favors preQ₁ affinity over preQ₀. Although structures of the B. subtilis aptamer also show that the preQ₁ methylamine is within hydrogen bonding distance of the phosphate backbone, this interaction is not observed in the T. tengcongensis aptamer bound to preQ₁ (this work) or preQ₀ (22). If this interaction were a means of discriminating the cyano group from the methylamine, the additional energetic difference favoring preQ₁ binding would be appreciable, adding another ∼3 kcal/mol (54). Further experiments will be necessary to establish the degree of conservation that this non-base interaction exhibits among family members and its role in preQ₁ affinity relative to preQ₀.

Despite the lower affinity of the T. tengcongensis aptamer for preQ₀ compared with preQ₁, the on-rates differ by only 12-fold, whereas the off-rates are comparable with preQ₀ dissociating 1.5-fold faster. Kinetic analysis of the QueF nitrile reductase (Fig. 1A) from B. subtilis showed a K_m of 237 μm with a k_cat of 0.69 min⁻¹, which are values comparable with the kinetic constants of other Q synthesis pathway enzymes (16). Assuming that these enzymes operate at steady-state levels and that K_m is a reasonable upper limit for substrate dissociation from the enzyme-substrate complex (55), then it is reasonable that preQ₀ concentrations are abundant in the cell at levels far exceeding its measured affinity in the context of the T. tengcongensis aptamer (K_D = 35.1 nm). As such, it appears plausible that the T. tengcongensis aptamer could respond to preQ₀ in gene regulation. This possibility is in agreement with the prior work of Breaker and colleagues (21), who recognized that the 10-fold affinity difference measured for preQ₁ versus preQ₀ in the context of the B. subtilis aptamer could not exclude preQ₀ as a physiologically relevant target.

Proposed Model of Translational Attenuation

The T. tengcongensis preQ₁ riboswitch represents an example of direct translational attenuation by the criteria that the aptamer is located fewer than 15 nucleotides upstream of the start codon (supplemental Fig. S1) and the sequestration of RBS sequences occurs within the aptamer domain (1). The preponderance of investigations on genetic regulation by riboswitches has focused on transcriptional control, thus making the T. tengcongensis aptamer a notable exception. Although the cellular concentrations of preQ₁ are not available in the literature, we can invoke the known K_m of 390 nm for the E. coli tRNA transglycosylase (56), which incorporates preQ₁ into tRNA and provides a reasonable upper limit for the metabolite's cellular concentration. Because the T. tengcongensis aptamer exhibits a K_D for preQ₁ of 2.05 nm (Table 3), it would appear that the aptamer senses preQ₁ at concentrations ∼200-fold lower than its steady-state consumption level. This seems plausible because the regulated gene product, COG1564, is a putative membrane protein invoked in preQ₁ transport (21, 57). Because GTP levels in E. coli are as high as 1.3–4.9 mm (58), the cell may need to be significantly depleted of GTP before it abandons de novo Q synthesis to undertake preQ₁ scavenging. With an off-rate of 1.53 × 10⁻⁴ s⁻¹, the half-life for dissociation of preQ₁ from the T. tengcongensis aptamer is 1.4 h. By contrast, the half-life for decay of the average E. coli message is 5 min (59), with individual half-lives ranging from seconds to as long as 1 h (59). This time frame suggests that once preQ₁ binds to the T. tengcongensis aptamer, the associated mRNA will undergo significant translational attenuation. However, translational attenuation of riboswitches such as the T. tengcongensis aptamer must also consider kinetic control. In E. coli, transcription and translation are closely coupled via the NusE-NusG protein interaction (32), which would require regulation by the T. tengcongensis aptamer to occur at preQ₁ levels far above its K_D. Such models require additional testing and validation but are worth considering in light of the different K_D values associated with members of the preQ₁ riboswitch family.

Differences in Ligand-induced Conformational Changes among PreQ₁ Aptamers from Different Genera

The preQ₁ riboswitch represents the first opportunity to compare metabolite-induced conformational changes by aptamers from three different organisms. Prior analysis of the B. subtilis and F. nucleatum preQ₁ riboswitches suggested that aptamer folding was dependent strictly on the presence of ligand (23, 25, 26). Our SAXS analysis of the F. nucleatum aptamer supports such a ligand-induced conformational change (Fig. 4), and the associated Kratky analysis suggests only partial folding in the absence of preQ₁ (supplemental Fig. S2D). This finding is consistent with other free-state analyses of these aptamers in solution that showed formation of a hairpin structure attached to a disordered 3′-tail (23, 26). By contrast, the T. tengcongensis aptamer is highly compact in the free state (R_g^P = 17.0 Å, r_max = 57 Å) and indicates little change in compactness in the presence of preQ₁ (R_g^P = 16.5, r_max = 51 Å). These results demonstrate that small riboswitch aptamers can exhibit highly compact free states in contrast to prior assumptions (60) and that variable degrees of ligand-dependent folding among riboswitches of the same class may be commonplace, suggesting that the mechanism of action for a given aptamer must be evaluated on a case-by-case basis (61).

Expectations for Molecular Level Conformational Features in the Free State

Crystal structures of the T. tengcongensis aptamer exhibited compact conformations in the bound and free states with dimensions that concur with those measured in solution (Table 2). However, efforts to fit observed and calculated scattering profiles (Fig. 4A) revealed that the preQ₁-bound state is a significantly better descriptor of the solution conformation (with a χ² of 1.7, indicating a reasonable fit (11)) as compared with the free-state structure with a χ² of 5.8, which is considered poor (Fig. 4A). The discrepancy between the preQ₁-free-state crystal structure and the solution data most likely originates from the constraints of the crystal lattice rather than incompleteness of the free-state model, which is missing nucleotide A13 (Fig. 2C). Our test calculations indicated that such a deletion has only a nominal effect on χ². In contrast, the free-state crystal structure fits better to the preQ₁-bound SAXS data than the free-state scattering data. This result is best explained by the similar crystal-packing environments of both crystal forms, which produced r_max values of ∼51 Å (Table 2). In solution, only the preQ₁-bound sample has an r_max of 51 ± 5 Å, whereas the free state showed an r_max of 57 ± 6 A. This suggests that crystal packing favors a more bound-state-like r_max, which may be the main source of the discrepancy in the free-state scattering profiles. An analogous result was observed previously for the SAM I riboswitch in which the SAM-bound state showed a χ² value of 1.6 between the crystal structure and solution data, whereas the free-state aptamer had a χ² value of 5.1. Like position A14 of the T. tengcongensis aptamer, position A46 of the SAM I aptamer moves 7.5 Å in the free state to occlude SAM access (supplemental Fig. S3, A versus B) (11).

Although the T. tengcongensis and SAM I free-state crystal structures are not adequate descriptors of solution scattering data, it is worth noting some general structural expectations for their free-state conformations. The observation that their ligand-binding pockets are blocked (Fig. 2D versus supplemental Fig. S3B) also has been observed for free-state crystal structures of the lysine riboswitch (9, 10) and the U65C mutant of the pbuE adenine riboswitch (62). Because such conformations are ill suited to ligand binding, it is most plausible that they are in equilibrium with open states in solution (11). The plausibility of this model came from the conformational ensemble analysis of the SAM-I riboswitch in which both open and closed conformations were required to deconvolute the free-state solution scattering profile, producing a χ² value of 0.83 (11). Members of the ensemble exhibited substantial changes in the orientation of helical segments, demonstrating folding differences beyond localized open and closed states. We envision a similar ensemble for the preQ₁ riboswitch.

Implications for Long Range Signaling between the Ligand Pocket and the RBS

Our mutagenesis results on various bases in and around the metabolite-binding pocket suggested that single functional group changes often exhibit deleterious binding effects incommensurate with those expected from the loss of 1–2 hydrogen bonds. Investigations of protein and RNA enzymes have shown that changes distant from the active site can influence catalytic function because they are part of an interaction network that limits dynamics and provide proper orientation of catalytic functional groups (63, 64).

It is conceivable that riboswitch aptamers are subject to control by similar interaction networks and that these help sample conformations conducive to ligand binding while maintaining a fold that couples ligand binding to distant regions of the riboswitch. This organization could serve as a basis to control accessibility to gene regulatory sequences, such as the RBS. We favor this interpretation for the T. tengcongensis preQ₁ riboswitch because a series of interconnected movements were implied by the free-state structure as a result of inserting base A14 into the ligand-binding pocket (Fig. 3A, arrows). Of special note is the greater accessibility to the RBS in the free-state structure and its apparent uncoupling from the aptamer core through the loss of base stacking. It remains to be seen whether this mode of interaction will be a general phenomenon for riboswitches that utilize direct translation attenuation to regulate gene expression.