Unwinding Single RNA Molecules Using Helicases Involved in Eukaryotic Translation Initiation

doi:10.1016/j.jmb.2006.06.016

J. Mol. Biol. (2006) 361, 327–335

Unwinding Single RNA Molecules Using Helicases Involved in Eukaryotic Translation Initiation Steven Marsden 1 , Maria Nardelli 1 , Patrick Linder 2 and John E. G. McCarthy 1 ⁎ 1

Manchester Interdisciplinary Biocentre, University of Manchester, 131 Princess Street, Manchester M1 7ND, UK 2

Département de Microbiologie et Medecine Moleculaire, Centre Médical Universitaire Université de Genève, 1211 Geneva 4, Switzerland

The small (40 S) subunit of the eukaryotic ribosome may have to scan more than 2000 nucleotides (>600 nm) from its 5′cap recruiting point on an mRNA molecule before initiating on a translation start codon. As with many other processes in living cells, including transcription, editing, mRNA splicing, pre-rRNA processing, RNA transport and RNA decay, scanning is facilitated by helicase activity. However, precise quantitative data on the molecular mechanism of scanning, including the roles of helicases, are lacking. Here, we describe a novel atomic force microscopy (AFM)-based procedure to examine the roles of two yeast helicases, eIF4A and Ded1, previously implicated in translation initiation by genetic and biochemical studies. Our results show that eIF4A, especially in the presence of its “cofactor” eIF4B, promotes ATP-dependent unwinding of localised secondary structure in long RNA molecules under tensional loading, albeit only at high protein:RNA ratios. Thus eIF4A can act to separate only a limited number of base-pairs, possibly via a steric unwinding mechanism. In contrast, Ded1 is more effective in reducing (by up to 50 pN at an AFM loading rate of 14 nNs−1) the force necessary to disrupt an RNA stem-loop, and thus shows significant kinetic competence to facilitate fast unwinding. These single molecule experiments indicate that Ded1 is likely to act as the more potent unwinding factor on natural mRNA substrates. © 2006 Elsevier Ltd. All rights reserved.

*Corresponding author

Keywords: protein–RNA interactions; translation initiation; helicases; atomic force microscopy

Introduction The small ribosomal subunit in both prokaryotes (30 S) and eukaryotes (40 S) is responsible for controlling base-pairing between the tRNA anticodon and each mRNA codon during protein synthesis.1 The challenge for the eukaryotic 40 S subunit is that, unlike its prokaryotic (30 S) counterpart, it does not locate directly to the position of the mRNA AUG codon where protein synthesis begins. The sequence-independent 5′→3′ scanning process manifests characteristics suggestive of the action of a molecular motor, but the mechanism involved is not understood.2,3 In order to be able to scan Abbreviations used: 5′UTR, 5′ untranslated region; AFM, atomic force microscopy; ss, single-stranded; ds, double-stranded. E-mail address of the corresponding author: [email protected]

successfully from the 5′cap to the start codon, the small ribosomal subunit needs to be able to overcome intramolecular secondary structures that would otherwise block progress along the RNA.4 Detailed studies in mammalian and fungal systems have demonstrated that the inhibitory influence of stem-loop structures in the 5′ untranslated regions (5′UTRs) of mRNAs is related to the stability of these structures.5,6 RNA helicases are proteins that can disrupt stemloop structures and hence there is an expectation that proteins of this type should be capable of facilitating 40 S binding and scanning. Most RNA helicases contain variations of a DEAD box motif along with six other highly conserved motifs.7 For example, the eukaryotic initiation factor eIF4A is a DEAD box RNA helicase that contains these seven core motifs together with only minimal flanking sequences. eIF4A folds into a “dumbbell” structure involving two globular domains joined by a linker. Upon ATP binding, eIF4A switches to a more

0022-2836/$ - see front matter © 2006 Elsevier Ltd. All rights reserved.

328 compact, “closed” conformation that shows greater affinity for single-stranded RNA and returns to the more open dumbbell conformation after ATP hydrolysis.8 Mammalian eIF4A has been shown to separate RNA duplexes in a bulk unwinding assay9 and is thought to act via a poorly processive translocative mechanism involving many association and dissociation events by undergoing cycles of conformational change. Mammalian eIF4B enhances the unwinding activity of eIF4A either by binding to single-stranded RNA regions and preventing re-annealing or by indirectly improving the processivity of eIF4A10 (although evidence for a stable interaction between eIF4A and eIF4B is lacking). Yeast eIF4A has previously been found to be inactive in standard oligonucleotide duplex unwinding assays when acting alone, and only when mammalian (and not yeast) eIF4B was added could weak helicase activity be observed.11 As such, it is possible that yeast eIF4A activity follows a different mechanism to that proposed for mammalian eIF4A. Ded1 is a larger DEAD box RNA helicase that possesses the same core motifs as eIF4A with the addition of flanking sequences that are suspected to account for an increased processivity along RNA. Ded1 has been suggested to operate via a non-processive “rolling” mechanism in which the helicase binds consecutively to singlestranded (ss) and then double-stranded (ds) RNA at a ss/dsRNA junction.12 An early model of the function of eIF4A proposes that this factor, by virtue of helicase activity promoted by eIF4B, facilitates the progress of the ribosomal complex along the 5′ UTR by supporting ATP-driven unwinding of mRNA structures. 13 However, more recent reports have highlighted the uncertainty concerning the true role of eIF4A in the mechanism of ribosomal scanning.8,9,14 Moreover, genetic (functional) interactions between yeast DED1 and translation initiation factor genes have suggested that Ded1 is involved in translation initiation.15,16 Indeed, Ded1 is an essential ATPdependent RNA helicase that manifests far higher specific ATPase activity than eIF4A.12 Overall, therefore, the respective roles of eIF4A and Ded1 in the scanning process in yeast remain ill-defined. A critical requirement of any helicase suspected to promote scanning is that it should be capable of facilitating unwinding of localised internal structures within long RNAs. Most importantly, this should be reflected in a reduction in the force that needs to be exerted by the 40 S ribosomal subunit in order that it can pass through the structured region. In other words, an analysis of the thermodynamic competence of a helicase to reduce the applied force necessary to achieve unwinding of localised structure in the context of a long RNA molecule represents an important measure of its capability to facilitate scanning. The use of bulk oligonucleotide duplex unwinding assays has not enabled us to answer a number of the key questions regarding the function(s) of helicases in translation initiation. Here, we report how we have developed, and

Single RNA Molecule Unwinding by Helicases

applied, a novel procedure for comparing the competence of the eIF4A and Ded1 helicases to unwind localised structure in long RNAs at the single molecule level. The procedure involves the use of atomic force microscopy (AFM) to stretch single RNA molecules tethered at their 5′ and 3′ ends, thus approximating to the situation in which a translocating biomolecular complex might apply force to open an intramolecular secondary structure.

Results We adopted a procedure for tethering RNA molecules between an AFM slide and the AFM cantilever tip that was particularly suited to the study of helicases (Figure 1(a)). Rather than using DNA “handles” to form DNA-RNA duplexes that hold the two ends of the molecule,17 we thiolated the 5′end of the RNA and incorporated biotinylated-adenine bases into its poly(A) tail. The presence of two-dimensionally distributed populations of immobilized thiol-RNA-biotin molecules on gold-coated AFM slides could be demonstrated using AFM scanning with a streptavidin-coated Si3Ni4 tip in either contact mode (Figure 1(b)) or frictional force mode (Figure 1(c)). Moreover, we could demonstrate the dependence of packing density on the applied concentration of thiolRNA-biotin molecules by incubating a series of slides with streptavidin-coated microspheres and scanning using a non-coated tip (Figure 1(d)–(g)). We next compared the activities of eIF4A, eIF4A: eIF4B and Ded1 in an 11 base-pair (mixed G:C/A:U) RNA duplex unwinding assay (Figure 2(a) and (b)) and an ATPase assay (Figure 2(c) and (d)) under the conditions to be used in AFM force-distance curve analyses. The short duplex used in the unwinding assay is not a good mimic of the mRNA substrates found in vivo but this assay is a useful indicator of helicase activity. The results revealed that, in the absence of any mechanical tension applied to the duplex, eIF4A and eIF4A:eIF4B are incapable of separating the strands of the 11 base-pair duplex under the given conditions, whereas Ded1 acts as a comparatively effective helicase (Figure 2(a) and (b)). Ded1 also manifested a specific ATPase rate that was approximately 40 times higher than that of eIF4A, and approximately 25 times higher than that of eIF4A:eIF4B (Figure 2(c) and (d)). These results confirmed the expected behaviour of our protein preparations in the standard bulk phase assays. With the components of our test system successfully prepared and tested using the standard assays, we applied the AFM-based procedure to the study of secondary structure unwinding in the context of a long RNA molecule, which is the predominant type of substrate that these helicases will normally encounter in the cell. Stretching experiments were performed using single-stranded RNAs based on the yeast GCN4 5′UTR. In order to record reversible force-curves, we set the ramp size to the contour length of the RNA (usually ∼250 nm)

Single RNA Molecule Unwinding by Helicases

329

Figure 1. Immobilisation of RNA for use in AFM force spectroscopy. (a) A diagram summarising the method used for suspending an RNA molecule between the AFM tip and a gold-coated microscope slide. (b) An AFM contact mode scan of the RNA construct on the gold surface. (c) A Frictional Force Mode (FFM) scan of the RNA construct on the gold surface. (d)–(g) 3D contact mode AFM scans of 40 nm diameter microspheres that are tethered to the gold slide using 0, 27, 540 and 13,500 pM of RNA construct, respectively. These 3D-scans were produced by scanning at 1 Hz in air, using an uncoated Si3Ni4 AFM tip. The gold surface was pre-treated with 50 μl of a bead-blocking solution before addition of the RNA construct.

and then moved the AFM cantilever closer to the surface (see Materials and Methods for definitions of AFM-related terms). This meant that the tip stayed close to the surface throughout the approach-retract cycle and thus had a greater chance of picking up a biotinylated RNA 3′ tail. Once short adhesion length reversible features appeared, we slowly retracted the tip in order to maximise the distance over which the RNA was pulled without exceeding its contour length. On average, the tip followed 20–30 approach and retract cycles before the biotin-streptavidin interaction was disrupted.

Further retraction of the tip eventually resulted in pull-off of the RNA causing a sudden drop to zero force and then a series of featureless force-curves. The RNA force-curves presented here were created using this method and the AFM tip was only relocated if reversible curves were not obtained within ∼100 consecutive approach-retract cycles. Curves for inclusion in the pool for quantitative analysis were selected on the basis of a set of objective rules dictated by the known parameters of the system (see Materials and Methods for rules and statistics).

330

Single RNA Molecule Unwinding by Helicases

Figure 2. Bulk phase analysis of RNA helicase unwinding and ATPase activities. (a) RNA unwinding activity of eIF4A/B. (b) RNA unwinding activity of Ded1 helicase. DsRNA, 70 fmol 32P-labeled RNA duplex; DP, dissociated products of 70 fmol 32P-labeled RNA duplex incubated at 94 °C for 3 min; ssRNA, 0.6 pmol 32P-labeled 11 nt RNA oligo; No ATP, helicase reaction set up as usual but with no ATP present. Labeled are the levels of migration for the RNA duplex (ds) and the small RNA oligo (ss). (c) Initial rate of ATP hydrolysis as the concentration of eIF4A (▴) and eIF4A/B ■) is increased. (d) Initial rate of ATP hydrolysis as the concentration of Ded1 is increased (▴).

In the absence of a major stem-loop structure, the force-distance curves generated by tip retraction and tip approach are closely matched, except for variable features during the early phase of retraction that correspond to non-specific interactions between the RNA (or the AFM tip) and the AFM slide goldcoated surface and/or to disruption of small secondary structures that form naturally in the GCN4 RNA (Figure 3(a)). We next incorporated stem-loop structures into the RNA. The first stemloop chosen was a compact, yet comparatively stable, stem-loop with 25 GC pairs in its stem. Previous work has shown that such a structure is capable of inhibiting translation initiation in Saccharomyces cerevisiae by at least 95%.6 Inclusion of this stem-loop in the GCN4 RNA sequence (Figure 3(b)) resulted in the reproducible appearance of a major discontinuity in the retraction curve, which shows rapid extension accompanied by a sharp reduction in force. The distance from the surface at which this occurred varied by up to 30 nm because of the variable position along the poly(A) tail of the biotin bound to the AFM tip. A smaller discontinuity was observed when a 15 base-pair stem-loop was introduced into the same position in the RNA (Figure 3(c)). In each such case, the discontinuity occurs because the force on the RNA increases until the hydrogen bonds of the stem-loop are broken, at which point the tension in the RNA drops as the extra nucleotides released are pulled taut, whereupon the force increases once more. This feature is not repeated during relaxation of the RNA because

there is not enough time for a stem-loop of this size and stability to refold during the approach phase under these loading rate conditions. Introduction of a helicase resulted in a reduction in the force measured at the peak of the discontinuity. An example is shown of the effect exerted by Ded1 (Figure 3(d)). Analysis of thousands of such force-distance curves performed under a range of conditions yielded plots of the estimated force required to induce the major discontinuity as a function of added protein concentration (Figure 4(a) and (b)). There was some unavoidable variation in the noise levels in the respective force-distance curves that was attributable to the variation in size and shape of different AFM tips and to automatic rescaling by the software as RNA molecules of slightly different lengths are pulled using the same retract distance. The ability of each DEAD-box protein to reduce the apparent force required to induce unwinding was strikingly different. While eIF4A worked maximally at a concentration of 0.15 μM to reduce the apparent force required for stem-loop opening (at 14 nNs−1) by 40 pN, Ded1 helicase required a concentration of only 50 nM to reduce this force by more than 50 pN. In further experiments (data not shown), we also found that increasing the amount of eIF4B to a twofold excess over eIF4A reversed the effect shown in Figure 4(a), consistent with an earlier observation that eIF4B can act to promote RNA annealing.18 Applying mechanical force to molecular bonds yields absolute estimates of the disruption force that

331

Single RNA Molecule Unwinding by Helicases

Figure 3. Single-molecule AFM force spectroscopy experiments showing the effect of RNA helicases on the force required to open a stem-loop. (a) Example of an AFM force-curve derived from the stretching of a single molecule of GCN4 5′UTR RNA (modified to contain no AUG codons) alone. (b) Example of an AFM force-curve representing stretching of an GCN4 RNA molecule containing the GC-rich (25 base-pair) stem-loop (inset). The discontinuity feature resulting from stem-loop opening is indicated by a small arrow. The approach curve (blue) runs from right to left and the retract curve (red) from left to right. (c) Example of an AFM force-curve representing stretching of a GCN4 RNA molecule containing the smaller (15 base-pair) stem-loop (inset). The discontinuity feature resulting from stem-loop opening is indicated by a small arrow. (d) Example of an AFM force-curve representing stretching of a GCN4 RNA molecule containing the GC-rich (25 base-pair) stem-loop in the presence of 0.4 μM Ded1.

are dependent on the loading rate.19 The forcedistance curves shown in Figure 3 were obtained using a loading rate of 14 nNs−1, which allowed measurement times within a reasonable time frame without generating too much variation in force estimates (see higher loading rate values in Figure 4(c)). However, the apparent force required for disruption of the stem-loop base-pairs estimated at this loading rate needs to be adjusted if we wish to estimate the correct force that would apply at other loading rates. For example, extrapolation of estimated force back to lower loading rates (Figure 4(c)), which generally apply in biological systems of this type,20,21 provides a guide to the correction necessary to obtain the force applicable at zero loading rate. Assuming that the slope of the calibration curve remains constant in the region approaching zero loading rate, the correction is equivalent to a difference of 115 pN. This correction to zero loading rate results in an estimated force of approximately 35 pN for disruption of the 25 basepair stem-loop structure. A rigorous analysis would not assume that this correction is directly transferable to experiments in which the unwinding force is reduced by helicases, but we have no reason to expect that the calibration is invalidated by the presence of these proteins. The corrected value compares favourably with an estimate made on the basis of a theoretical model for mechanical disruption based on an extended Freely Jointed Chain model that also takes into account the minimum free energy of bond dissociation during the

unfolding pathway (40 pN).22 It also compares well with the force estimate for the unwinding of a 14 base-pair stem-loop using laser tweezers under near-equilibrium (zero-loading rate) conditions (25 pN).23 Most importantly, and irrespective of the absolute value for the disruption force, which will in any case vary somewhat according to which technique is used to measure it, the key finding is that Ded1 is a more effective helicase in this context than eIF4A (Figure 4(a)).

Discussion We conclude that the mechanochemical approach described here provides new insight into the distinct capabilities of the eIF4A and Ded1 helicases. By using the AFM-based procedure described here, we can apply tensional loading conditions that in principle approximate to the situation where a processive biomolecular complex (in this case a scanning 43 S pre-initiation complex) bears down on a localised internal RNA structure, and thus determine the relative capabilities of different helicases to facilitate localised unwinding in a long RNA molecule. Our results demonstrate that eIF4A, even in the presence of eIF4B, is a less effective facilitator of RNA stem-loop unwinding than Ded1, and thus prompt us to reconsider the likely roles of these two proteins in translation initiation. The fact that yeast eIF4A unwinding activity saturates well before the 25 bp stem-loop is fully unwound (as

332

Single RNA Molecule Unwinding by Helicases

Figure 4. Plots of helicase concentration against unwinding force. (a) Stem-loop unwinding forces were measured over a range of concentrations of eIF4A/B (●) and Ded1 (▴). The control curve shows data collected using BSA instead of a helicase (■). Standard error bars are included. (b) The effect of increasing RNA helicase concentration on the opening force of the stem-loop in the presence of ATP plotted along with negative controls where no ATP was added. Stem-loop unwinding forces were measured over a range of concentrations of eIF4A/B (●) and Ded1 (▴) in the presence of ATP (5 mM). Control experiments lacking ATP were also performed for eIF4A/B (■) and Ded1 (♦). RNA was bound to the gold slide as previously described and AFM force spectroscopy was carried out in 50 μl of AFM pulling buffer (0.5 unit of RNasin, 10 mM MgCl2, 250 mM NaCl, 10 mM Tris-HCl in DEPC-treated water) at ∼1 Hz using a trigger force of 0.5 nN, a surface delay of 400 ms, a retract velocity of 1 μm/s and a ramp length of ∼250 nm with a streptavidin-coated S3N4 AFM cantilever of spring constant 0.02 N/m. (c) Plot of estimated mean opening force for the 25 bp stem-loop as a function of loading rate. These data points were fitted using the software Curve Expert 1.3 according to the quadratic equation y = 35.25 + 8.39x−0.24x2. (d) A theoretical retraction force curve for the stretching of the GCS4 L1-RNA transcript (with a 65 nt poly(A) tail). This trace is adapted from data generated using the online “RNA pulling server” at http://www. bioserv.mps.ohio-state.edu/rna. 22 The model assumes a temperature of 37 °C, 1 M NaCl, and a nucleotide length of 0.334 nm. The predicted ∼40 pN GC-rich stem-loop opening feature is labelled with an arrow.

shown by the data in Figure 4(a)) suggests that this helicase operates via a (comparatively slow) steric mechanism. This corresponds to a model in which eIF4A operates by binding preferentially to singlestranded regions of the 5′UTR, whereby the presence of the helicase at the base of a stem-loop structure enables ATP-induced conformational changes in eIF4A to cause partial unwinding (Figure 5(a); compare Lorsch & Herschlag). eIF4B promotes eIF4A function primarily by enhancing the affinity of eIF4A for single-stranded RNA.18 The role of ATP-induced conformational changes in eIF4A is supported by an earlier study of eIF4A based on limited proteolysis.8 Ded1, on the other hand, can facilitate unwinding of almost the entire stem-loop at a lower concentration than is required for maximal eIF4A activity. Moreover, our data indicate that Ded1 shows a greater kinetic competence (in the presence of ATP), meaning that it can bind to newly exposed single-stranded RNA and contribute to unwinding at rates comparable to the AFM-induced unzipping of the stem-loop. This greater kinetic competence relative to eIF4A might also reflect the

involvement of a degree of processivity in the Ded1 mechanism and so it is likely to follow a lowprocessivity translocative model for RNA unwinding of the type suggested previously12 (see mechanism in Figure 5(b)), but this cannot be deduced from the AFM data alone. Finally, these qualitative and quantitative mechanistic differences between the two types of helicase now suggest an explanation for the previous observation 14 that Ded1, but not eIF4A/eIF4B, preferentially promotes the translation of mRNAs bearing longer, more structured 5′ UTRs in vivo. Why should the cell utilize more than one helicase to facilitate translation initiation? One potential explanation of this would be in terms of distinct areas of function, as illustrated by the model depicted in Figure 5(c). By virtue of its association with the eIF4G component of the cap-binding complex, eIF4A activity is likely to be focused on the 5′region of the mRNA, where it can help enable efficient 43 S recruitment. In line with this interpretation, Bordeleau and colleagues24 have concluded that pateamine A, a newly discovered

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Single RNA Molecule Unwinding by Helicases

Figure 5. Working models of RNA unwinding by the helicases. (a) In the steric unwinding model, the affinity and kinetics of interaction between the helicase (purple) and ssRNA determine how effectively unwinding is facilitated. Conformational changes (square to triangle) in the helicase associated with ATP binding (hydrolysis) enhance affinity for ssRNA. A helicase with a comparatively low kon will be less kinetically competent to promote unwinding. (b) In a processive translocative model, ATP binding and hydrolysis drive movement of the helicase (brown circle and oval) along the RNA, thus dynamically promoting unwinding. (c) The two types of helicase may contribute in different ways to the initiation process. The primary domain of activity of eIF4A, whose mechanism approximates to the steric model (a), may be focused in the vicinity of the 5′cap by virtue of this protein's association with the eIF4F complex (the components of this complex, eIF4E, eIF4G and eIF4B, are shown in the cartoon; eIF4A is represented by purple triangles). In contrast, the more dynamic mode of action of Ded1, which may follow a combination of mechanisms (a) and (b), makes this helicase more suited to promoting rapid unwinding of internal structures.

specific ligand of eIF4A that inhibits translation initiation, acts by affecting 43 S recruitment to mRNA. The more dynamic helicase activity of Ded1, in contrast, is expected to be more suited to facilitating structural unwinding under load within the kinetic framework of the scanning process once the 43 S pre-initiation complex ventures beyond the cap-proximal region of the mRNA.

Materials and Methods RNA The L1 GCN4 leader sequence from S. cerevisiae (with all AUG codons removed, as described)20 was used as the basis for this work. Stem-loop containing RNA segments

were encoded by fragments derived from plasmids described earlier.25 Transcriptions were carried out using an RNAmaxx kit (Stratagene) and 1μg linearised plasmid to yield ∼100 μg RNA in a final volume of 25 μl. Two units of DNase (Promega) per μg of DNA were added at the end of the transcription reaction, followed by a further incubation at 37 °C for 30 min. The RNA was then phenol-extracted, ethanol-precipitated, resuspended in 50 μl of DEPC-treated water and run on a ProbeQuant G-50 microcolumn (Amersham Bioscience) to remove unincorporated nucleotides. 5′Thiol-incorporation was achieved using a ”5‘end-tag” kit (Vector Laboratories). First the transcripts were dephosphorylated in 1× universal buffer, 0.5 unit/μl RNase inhibitors and alkaline phosphatase in a final volume of 10 μl DEPC-treated water, incubated at 37 °C for 30 min. The thiol group was then attached immediately by adding 0.5 unit/μl RNase inhibitors, 1× universal buffer, 1 mM ATPγS and Polynucleotide kinase in a final volume of 20 μl DEPC-treated water, incubated at 37 °C for 30 min. The RNA was then phenol-extracted and ethanol-precipitated. Polyadenylation was used to incorporate biotinylated nucleotides at the 3′end. RNA was incubated with 1× PAP buffer (20 mM Tris-HCl (pH 7.0), 50 mM KCl, 0.7 mM MnCl2, 0.2 mM EDTA, 100 μg/ml acetylated BSA, 10% (v/v) glycerol (USB)), 1.3 unit/μl RNasin, 0.13 mM bio-17-ATP (Sigma), 3.3 mM ATP and 20 units/μl poly(A) polymerase made up to a final volume of 15 μl with DEPC-treated water. The reaction was incubated at 37 °C for 30 min and purified on ProbeQuant G-50 microcolumns and 30 μl of eluate was recovered. Atomic force microscopy AFM and FFM scans were produced simultaneously using a streptavidin-coated Si3Ni4 AFM tip (with a spring constant of 0.02 N/m), scanning at 1 Hz in 50 μl of imaging solution (250 mM NaCl, 10 mM MgCl2, 10 mM Tris-HCl (pH 7) and 0.1 unit of RNasin). RNA was bound to the gold surface in 10 μl of binding buffer (1.3 unit RNasin, 1× BSA, 10 mM MgCl2, 250 mM NaCl, 10 mM Tris-HCl in DEPC-treated water) at room temperature for 20 min and washed with 200 μl imaging solution. 3D contact mode scans were produced by scanning at 1 Hz in air, using an uncoated Si3Ni4 AFM tip. The gold surface was pre-treated with 50 μl of a bead-blocking solution before addition of the RNA construct. AFM force spectroscopy was carried out in 50 μl of AFM pulling buffer (0.5 unit of RNasin, 10 mM MgCl2, 250 mM NaCl, 10 mM TrisHCl in DEPC-treated H2O) at ∼1 Hz using a trigger force (the positive force that must be reached in the tip approach phase before retraction from the surface occurs) of 0.5 nN, a surface delay of 400 ms, a retract velocity of 1 μm/s and a ramp length of ∼250 nm with a streptavidin-coated S3N4 AFM cantilever of spring constant 0.02 N/m. We applied a set of rules to identify force-distance curves that corresponded to the expected behaviour of single RNA molecules that was not unduly distorted by artefactual interactions. In order to record reversible force-curves, we set the ramp size (the maximum vertical displacement of an AFM tip above the sample surface) to the contour length (the end-to-end distance of a polymer once it is fully stretched) of the RNA (usually ∼250 nm) and then moved the AFM cantilever closer to the surface In brief, we included in our analysis only those thiolRNA-biotin force curves that were reversible, that manifested an initially gradual slope, and that reached an adhesion length (the length to which a bond,

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Single RNA Molecule Unwinding by Helicases

Table 1. Force-distance curve statistics GC-rich stem-loop statistics

Sample Th-GC-rich stemloop-RNA-Biotin +eIF4A+ATP +eIF4A+B+ATP +Ded1+ATP a

Opening force (pN)a % Frequency Mean SD Median Mode 71.9

151.3

29.75

150

150

51.7 25.6 8.5

135.2 114.1 94.6

32.98 34.12 23.3

140 110 90

100 110 80

Values not corrected for loading rate.

interaction or polymer can be stretched before the molecule is pulled off the tip or surface during AFM force spectroscopy measurements) of at least 250 nm and a force of at least 180 pN. The GC-rich stem-loop opening feature on each of these selected curves was identified as a single discontinuity found in the last 50 nm of the retract curve. We recorded the initial force at which each discontinuity began. Curves with a single discontinuity force larger than 250 pN in this region were discounted since such extraordinarily large forces could only be due either to stabilisation of the stem-loop by protein binding or to pulling of aligned RNA aggregates. In this way, we determined the percentage of the Th-RNA-biotin forcecurves that manifested the GC-rich opening feature. The initial force values from these selected curves were used as the basis for estimation of average disruption forces. Table 1 shows a summary of the statistics compiled for the percentage of RNAs with the GC-rich stem-loop and the opening force for RNAs with this feature in the absence of any helicase, and in the presence of excess ATP along with 0.4 μM eIF4A, 0.4 μM eIF4A/B or 0.4 μM Ded1 (>1000 RNA force-curves were used in each case). The data for the dependence of the GC-rich opening force on the concentration of eIF4A/B or Ded1 were generated using this manual counting method, where at least a thousand curves were used for each helicase concentration investigated. Supplementary Figure 1 shows further examples of force-distance curves, illustrating typical variations in the magnitude of the discontinuity. Duplex unwinding assay Unwinding reactions followed a previously described procedure26 and were set up with 0.4 μM RNA helicase, 1 mM ATP, 10 mM Tris-HCl, 250 mM NaCl, 10 mM MgCl2, and one unit of RNasin, started by the addition of 70 fmol of an 32P-labeled RNA partial duplex (comprising an 11 bp duplex region of sequence 5′-GACCGUAAAGC-3′ and a 14 nt 3′-overhang of sequence 5′-ACGCAAAAACAAAA3′), incubated at 35 °C and stopped with 5 μl of helicase stop buffer (50% (v/v) glycerol, 2% (w/v) SDS, 5 mM EDTA, 0.1 unit/μl RNasin, 0.01% (w/v) Bromophenol blue) after 0, 0.5, 1, 1.5, 2, 4, 8, 15 or 30 min. ATPase assay ATPase reactions27 were set up with 4 pmol (859 ng) RNA in 65 mM Tris-HCl, 1.3 μM MgCl2, 1.3 μM DTT, 1 μg Taxol, 0.21 mM NADH, 1 mM Mg.ATP, and 3 mM phosphoenolpyruvate, and were started by the addition of 7.4 units of pyruvate kinase, 10.5 units of lactate dehydrogenase and RNA helicase. A340nm was measured every 5 s for 300 s and plotted against time. Rates were

derived from the initial slope for A340 change over time at each helicase concentration. ATP hydrolysis was first converted into μM s−1 by dividing by 0.00622 μM−1 and then into nmol s−1.

Acknowledgements We thank the Biotechnology and Biological Sciences Research Council (BBSRC), the Engineering and Physical Sciences Research Council (EPSRC), the Royal Society and the Wolfson Foundation for supporting this work.

Supplementary Data Supplementary data associated with this article can be found, in the online version, at doi:10.1016/ j.jmb.2006.06.016

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Edited by D. E. Draper (Received 30 April 2006; received in revised form 7 June 2006; accepted 9 June 2006) Available online 27 June 2006