Helicase-mediated changes in RNA structure at the single-molecule level

Abstract

RNA helicases are a diverse group of RNA-dependent ATPases known to play a large number of biological roles inside the cell, such as RNA unwinding, remodeling, export and degradation. Understanding how helicases mediate changes in RNA structure is therefore of fundamental interest. The advent of single-molecule spectroscopic techniques has unveiled with unprecedented detail the interplay of RNA helicases with their substrates. In this review, we describe the characterization of helicase-RNA interactions by single-molecule approaches. State-of-the-art techniques are presented, followed by a discussion of recent advancements in this exciting field.

Keywords: AFM; DEAD-box; FRET; PIFE; RNA folding; RNA helicase; optical tweezers; single-molecule spectroscopy.

Figure 1. Helicase-induced RNA hairpin unfolding using AFM force spectroscopy. (A) RNA immobilization involves tethering the 5′-end to the gold-coated surface (“Au”) and attaching the biotinylated 3′-end (“B”) to a streptavidin-coated (“S”) silicon nitride (Si₃Ni₄) tip. Force-extension curves were recorded in the presence and absence of eIF4A or Ded1, represented as “HEL.” (B and C) Representative force-extension plots representing stretching of a single RNA molecule containing a GC-rich stem-loop. (C) Stretching of the same RNA in the presence of 400 nM Ded1. The pull curve (red) runs from left to right and the approach curve (blue) from right to left. The arrow indicates the force and extension when the hairpin unfolds. In the presence of Ded1, the hairpin unfolds at lower force. Adapted from reference .

Figure 2. Revealing NS3 translocation and unwinding mechanism by optical tweezers. (A) A 60 bp RNA hairpin is flanked by two RNA/DNA duplexes. The bottom duplex (599 bp) is attached to a bead via biotin-streptavidin linkage (“B,” “S”). The top duplex (535 bp) is attached to an anti-digoxigenin-coated bead via digoxigenin (“A,” “D”). NS3-mediated (“HEL”) duplex unwinding is followed upon application of external force. (B) Unwinding experimental steps: mechanical folding and unfolding of substrate in the absence of NS3 shows a transition at 20.4 pN, the force required for hairpin unwinding (green), the presence of NS3 decreases the force required for duplex unwinding (red), force is brought to 30 pN to fully extend the nucleic acid strand (blue). The RNA hairpin is allowed to refold at 2 pN force (yellow). (C) Extension of bead separation during RNA unwinding by NS3 reveals discrete steps and pausing. (D) A pairwise distance histogram of the unwinding trace shown in (C) is later subjected to Fourrier analysis to determine the apparent unwinding step size. (E) Unified model for NS3 helicase activity. Nucleic acid substrate binding is followed by ATP-dependent destabilization duplex by the translocator domain (blue ellipse). The RNA hairpin is unwound by helix-opening domain (orange circle) in fast ATP-dependent 1 bp-substeps adding up to steps of 3.6 ± 1.3 bp, unwinding 7–13 basepairs per apparent step depending on the GC content.^,,, Subsequently, the translocator moves forward to start a new catalytic cycle. At this stage, GC-rich sequences increase the probability of helicase dissociation. Adapted from references and .

Figure 3. Ded1-mediated RNA remodeling monitored by smFRET. (A) Experimental design and results. The Cy3-labeled RNA strand (light gray; Cy3, green circle) forms a thermodynamically stable duplex with the immobilization strand (white) that is surface-tethered via a biotin-streptavidin linkage (“B,” “S”), while the Cy5-labeled strand (dark gray; Cy5, red circle) is in solution. Strand exchange in the presence of Ded1 (“HEL”) and in the absence of ATP proceeds via a tripartite intermediate characterized by a sudden increase in Cy5 emission through FRET, followed by the formation of the thermodynamically less stable duplex accompanied by a complete loss of fluorophore emission (upper pathway). In the presence of both Ded1 and ATP, strand exchange involves complete disassembly of the more stable duplex and subsequent formation of the less stable one (lower pathway). Passivation of the quartz slide is achieved through coating with polyethylene glycol to prevent non-specific binding of the protein to the slide. (B and C) Representative fluorophore emission and FRET time traces. Cy3 and Cy5 emission over time (green and red traces) displays FRET-typical anti-correlated behavior (upper plots). FRET over time reveals the formation of the tripartite intermediate, as indicated by a sudden burst of FRET (lower plots, black arrows), followed either by dissociation of the Cy5-labeled strand (B) or strand exchange and complete absence of fluorophore emission (C, red arrow). Figure modified from reference .

Figure 4. Characterizing NPH-II-catalyzed RNA duplex unwinding. (A) Experimental design and results. A fluorophore-labeled 19-bp RNA duplex (Cy3, green circle; Cy5, red circle) with a 24-nt 3′ extension was immobilized on the PEG-passivated quartz slide via a biotin-streptavidin linkage (“B,” “S”). The initial high FRET value of 0.85 is diminished in response to NPH-II binding (“HEL”) and fluctuates between two discrete low FRET values, indicating an increased inter-dye distance. Addition of ATP triggers duplex unwinding ultimately leading to complete loss of emission upon strand separation. (B) Representative smFRET trajectory (1 nM NPH-II, no ATP) showing transition between the helicase-unbound state (FRET 0.85, highlighted in orange), and the helicase-bound states (FRET 0.15 and 0.33, highlighted in green and yellow, respectively). (C–H) Averaged FRET histograms, each built from over 100 individual FRET time traces. Imaging conditions: (C) only RNA, (D) RNA and 100 nM NPH-II, (E) 100 nM NPH-II, 3.5 mM ATP, (F) 100 nM NPH-II, 3.5 mM 'ADP-BeF_x (a ground-state analog), (G) 100 nM NPH-II, 3.5 mM ADP-AlF_x (a transition state analog), (H) 100 nM NPH-II, 3.5 mM ADP. (I) Basic model for unwinding initiation by NPH-II relying on altered substrate affinities along the ATP hydrolysis cycle. Without nucleotide, NPH-II binds both ssRNA and dsRNA and the NPH-II-ssRNA complex readily alternates between two conformations. ATP binding impedes dissociation from ssRNA and changes the kinetics of bound-state transitions. In the ATP transition state, NPH-II no longer binds to dsRNA and interconversion kinetics change again. The helicase associates with dsRNA upon ATP hydrolysis and phosphate dissociation. Different shapes mark the different conformational states if NPH-II traversed during unwinding initiation. Figure adapted from reference .

Figure 5. Mss116-mediated group II intron folding using smFRET.^, (A) Experimental design. The fluorophore-labeled D135 ribozyme (Cy3, green cycle; Cy5, red cycle) is immobilized on a PEG-coated quartz slide via a biotin-streptavidin linkage (“B,” “S”). Structural interconversion under different folding conditions is monitored by following FRET efficiency over time. (B–F) Averaged FRET histograms, each built from over 100 single-molecule time traces. Imaging conditions: (B) 8 mM Mg²⁺, 500 mM K⁺. Three FRET distributions are observed, termed “intermediate” (“I”), “folded” (“F”) and “native” (“N”) based on earlier results.^, (C)Eight mM Mg²⁺, 100 mM K⁺. Only the “I” FRET state is observed at near-physiological conditions. (D) Eight mM Mg²⁺, 100 mM K⁺, 25 nM Mss116, 1 mM ATP. Addition of Mss116 and ATP shifts the distribution of FRET states toward the folded intermediate and the native state. (E and F) Eight mM Mg²⁺, 100 mM K⁺, 25 nM Mss116 (and 1 mM AMPPNP). Effect of ATP hydrolysis on D135 folding. Prevalence of the native state is lowered in the absence of ATP (E) and in the presence of non-hydrolyzable AMPPNP (F). (G) Percentage of dynamic molecules at different imaging conditions. Red, 500 mM K⁺, 8 mM Mg²⁺; green, 8 mM Mg²⁺, 100 mM K⁺; blue, 8 mM Mg²⁺, 100 mM K⁺, 25 nM Mss116, 1 mM ATP; yellow, 8 mM Mg²⁺, 100 mM K⁺, 25 nM Mss116; purple, 8 mM Mg²⁺, 100 mM K⁺, 25 nM Mss116, 1 mM AMPPNP. (H) Proposed model of Mss116-mediated group II intron folding. D135 interconverts between four conformations referred to as “unfolded” (“U”), “intermediate” (“I”), “folded” (“F”) and “native” (“N”). Mss116 promotes the transition from U to I, even in the absence of ATP. It further catalyzes ATP-dependent conversion from F to N. Figure modified from reference .

Figure 6. Probing RIG-I translocation on dsRNA using PIFE. (A and B) Experimental design. A 25/40 bp dsRNA with blunt ends (A) or a 20–50 bp dsRNA with a 66–36 nt ssRNA overhang and a 5′ triphosphate (“PiPiPi”). RNA construct (B) is labeled with a single DY547 fluorophore (green circle) and immobilized on a PEG-passivated surface via biotin-neutravidin (“B,” “N”). (C) Depiction of three modular RIG-I variants used in this study. RIGh consists of the central DExH-box ATPase domain and a C-terminal regulatory domain (“RD”). wtRIG additionally has two N-terminal caspase activation and recruitment domains (“CARD”). In svRIG, one of the CARDs is non-functional. (D) Representative time trajectory recorded in the presence of RIGh and the blunt end RNA substrate shown in (A). Helicase binding is accompanied by a sudden increase of fluorophore emission. (E–G) Dwell-time analyses for time traces recorded in the presence of RIGh (E), wtRIG (F) and svRIG (G) for 25-bp and 40-bp blunt end dsRNA shown in (A). RIGh and svRIG translocate faster along dsRNA than wtRIG. In all cases, the average time required for end-to-end translocation increases with the substrate length. (H) RIG-I translocation on dsRNA in the presence of 5′-triphosphate, average time spent in the bound state vs. duplex length. (I) Proposed model for pathogen-associated molecular pattern (“PAMP”) signal integration by RIG-I. Binding of the RIG-I regulatory domain (pink) to RNA 5′ triphosphates induces RIG-I dimerization as described previously. This triggers the translocase domain (blue), followed by translocation along the dsRNA substrate (red arrow) and induction of a CARD signaling conformation (gray). Figure modified from reference .