The Structure and Folding of Branched RNA Analyzed by Fluorescence Resonance Energy Transfer

C H A P T E R

E I G H T

The Structure and Folding of Branched RNA Analyzed by Fluorescence Resonance Energy Transfer David M. J. Lilley Contents 162 163 167 170 171 176 179 182 182

1. Theory of FRET 2. The Possible Effects of Fluorophore Orientation 3. Choice of Fluorophores 4. Construction of Fluorophore-Labeled RNA Species 5. Steady-State Measurements of FRET 6. Time-Resolved Measurements of FRET 7. Single-Molecule FRET Acknowledgments References

Abstract Fluorescence resonance energy transfer (FRET) is a spectroscopic means of obtaining distance information over a range up to
80 A˚ in solution. It is based on the dipolar coupling between the electronic transition moments of a donor and acceptor fluorophore attached at known positions on the RNA species of interest. It can be applied in ensembles of molecules, either by steady-state fluorescence or by lifetime measurements, but it is also very appropriate for single-molecule studies. In addition to the provision of distance information, recent studies have emphasized the orientation dependence of energy transfer.

Fluorescence (or Fo¨rster) resonance energy transfer (FRET) (Fo¨rster, 1948; Perrin, 1932) provides a spectroscopic way of estimating distances over a size range that is appropriate for biological macromolecules. It is based upon fluorescence, one of the most sensitive spectroscopic methods. Fluorescence is the emission of light from an excited molecule, having lost Cancer Research UK Nucleic Acid Structure Research Group, MSI/WTB Complex, The University of Dundee, Dundee, United Kingdom Methods in Enzymology, Volume 469 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)69008-X

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2009 Elsevier Inc. All rights reserved.

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vibrational energy in the excited state. Thus, the fluorescent photon is of longer wavelength than the photon that excited the molecule, the wellknown Stokes shift. Molecules interact with the electric component of light because of the change in electronic distribution between the ground state and an excited state, measured by the transition dipole moment vector. ^ can be written in Thus, the vector for a transition from states m to n (d) bra.ket notation as: d^ ¼ hCn jℜjCm i

ð8:1Þ

where Cm and Cn are the wavefunctions of the initial and final states, respectively, and ℜ is the dipole moment operator. The transition dipoles of two different fluorescent molecules may interact together in a dipolar process that leads to transfer of energy from one (the donor) to the other (the acceptor). This resonance energy transfer is strongly dependent upon the physical separation between the two. By tethering two smallmolecule fluorescent probes (fluorophores) to a biomolecule of interest at known positions, we can monitor the distance between these two points. For other reviews, see Clegg (1992, 2002), Lakowicz (1999), Walter and Burke (2000). Although FRET has been used in biochemical studies for half a century or more (Bennett, 1964; Cantor and Pechukas, 1971; Dale and Eisinger, 1976; Stryer and Haugland, 1967), its full exploitation in the study of nucleic acid structure and folding required the ability to synthesize oligonucleotides chemically and to attach fluorophores at known positions, and that has only been possible in the last 20 years. One of the first such studies of branched nucleic acids was the analysis of the structure of the four-way (Holliday) DNA junction, in which junctions with two arms terminally labeled with fluorescein and tetramethylrhodamine were studied in the steady state (Murchie et al., 1989). Since that time, FRET has been extensively used to study the structure and folding of branched nucleic acids, or junctions. Here, we define helical junctions as structures in which a number of helical segments are connected by the covalent continuity of strands shared between them (Lilley, 2000). There may or may not be additional unpaired nucleotides present at the positions where the strands exchange between the helices. These are named according to the IUB nomenclature (Lilley et al., 1995). The most common junctions found in natural RNA are three- and four-way junctions. Such junctions can act as important architectural elements in large RNA species, or as key folding motifs in small autonomously folding RNAs. We may broaden this definition to include a duplex containing a central structural feature that leads to a discontinuity in the axis; this could be regarded as a two-way helical junction (Gohlke et al., 1994). The kink-turn (k-turn) (Klein et al., 2001) is a good example of a two-way junction.

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In a typical FRET experiment involving branched RNA or DNA, the nucleic acid would be labeled with donor and acceptor fluorophores, covalently attached at different locations. Analysis of FRET between the fluorophores then provides an estimate of the distance between them. Estimates of the absolute distance could provide input into molecular modeling to determine the structure, but the difficulties of getting accurate distances from FRET information are discussed below. Nevertheless, even relative information can be valuable. A series of vectors within the structure might be compared. For example, the three end-to-end distances of a threeway junction, or the six end-to-end distances of a four-way junction might be analyzed. Further information could be derived by systematic variation of the length of the helix to which a given fluorophore is attached. In some experiments the absolute value of FRET efficiency (or distance) may be less important than how it changes, for example, during folding induced by metal ion concentration or protein binding. FRET experiments on RNA are generally carried out in one of the three main ways. The simplest is by steady-state fluorimetry, where a solution of fluorophore-labeled RNA is excited at the wavelength of the donor, and then the intensity of the emitted light is scanned over the range of donor and acceptor emission. It requires relatively simple equipment, and unsophisticated data processing. FRET efficiency can be determined from the reduced fluorescence of the donor in the presence of the acceptor, or the enhanced emission of the acceptor. However, steady-state data are an average over the ensemble of molecules present, and no information is obtained about subpopulations of species within the ensemble. The second approach is to analyze the excitedstate lifetime of the donor. This becomes shortened in the presence of an acceptor due to the transfer of excitation energy by FRET, and so provides a measure of the efficiency of the process. Like the steady-state measurements, this is performed on a solution of donor–acceptor-labeled RNA. But the timedependence of the decay of donor molecules in the excited state can be fitted to models in which multiple populations of species have distinct fluorescent lifetimes, and the distributions of the different species can be estimated. Time-resolved fluorimetry requires a higher level of sophistication in terms of equipment and data processing. Lastly, the high sensitivity of fluorescence means that FRET experiments can be performed on single RNA molecules, either tethered to a glass slide in some manner, or freely diffusing in solution. Single-molecule FRET offers a number of advantages over ensemble methods. The ability to examine one molecule at a time allows us to divide the population into its constituent species, reveal heterogeneity within seemingly identical molecules, study molecules within impure samples or even cell extracts, and study kinetic processes that cannot be synchronized. There are a variety of ways to carry out single-molecule FRET experiments, all of which are based on microscopy and highly sensitive detection methods. Most laboratories have assembled their own equipment to perform this.

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1. Theory of FRET FRET involves a resonance between singlet–singlet electronic transitions of the donor and acceptor fluorophores, arising from the dipolar coupling between the emission transition dipole of the donor and the absorption transition dipole of the acceptor. This leads to a transfer of excitation energy from the donor to the acceptor. The process occurs within a region much smaller than the wavelength of the light (the near field region), and therefore does not involve the transfer of real photons. FRET can be observed in a variety of ways, including a reduction in the fluorescent quantum yield of the donor, a corresponding shortening of the donor excited-state lifetime, and an increased fluorescent emission from the acceptor (if fluorescent). Either a classical or quantum mechanical analysis shows that the rate of the energy transfer process depends on the inverse sixth power of the distance between the two fluorophores (Fo¨rster, 1948). This is the basis of the use of the technique to provide structural information. In the laboratory the efficiency of the process (EFRET) is normally determined. This can be defined in a variety of ways, and is the proportion of donor excitation events that lead to excitation of the acceptor by dipolar coupling—a quantum yield of FRET: "
6 #1 1  tDA rDA kFRET EFRET ¼ ¼ 1þ ¼X ð8:2Þ tD R0 k i i where tDA and tD are the fluorescent lifetime in the presence and absence of the acceptor, respectively, rDA is the separation between the donor and acceptor. This results in the dependence on distance shown graphically in Fig. 8.1. EFRET can also be defined in terms of rates, where kFRET is the rate constant for energy transfer and ki are those for all the mechanisms of deactivating the donor excited state (including fluorescent emission from the donor, quenching, intersystems crossing, etc.). R0 is the characteristic Fo¨rster length for a given donor–acceptor pair of fluorophores, given by: 0:529  FD k2 JðlÞ ð8:3Þ Nn4 where the units of R0 and the wavelength l are in cm. FD is the fluorescent quantum yield of the donor in the absence of the acceptor and N is the Avogadro number. k is a parameter that depends on the relative orientation of the donor and acceptor transition moments, discussed in the following section. n is the refractive index of the medium in which the electric fields of the transition dipoles extend; a value of 1.33 is appropriate for the aqueous medium of biological macromolecules. J(l) is the normalized spectral overlap between donor emission and acceptor absorption, given by: R06 ¼

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1

E FRET

0.8

0.6

0.4

0.2

0

0

0.5

1 r DA/R 0

1.5

2

Figure 8.1 The dependence of FRET efficiency (EFRET) as a function of the separation of donor and acceptor fluorophores (rDA) normalized to the Fo¨rster distance R0 (Eq. (8.2)), assuming a constant value of R0.

ð1 JðlÞ ¼

0

fD ðlÞeA ðlÞl4 dl ð1 fD ðlÞ dl

ð8:4Þ

0

where fD is the spectral shape function for the donor emission and eA is that for acceptor absorption (in M 1 cm 1). From equation (8.2) it can be seen that when rDA ¼ R0, the efficiency of FRET is 0.5. R0 values are frequently calculated on the basis of an assumption (stated or otherwise) that k2 ¼ 2/3. Indeed, this can be written as R0(2/3).

2. The Possible Effects of Fluorophore Orientation FRET arises from the dipolar coupling between the oscillating transition dipoles of the donor and acceptor fluorophores. The magnitude of the interaction depends on both the distance between them and their relative

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orientation. The rate at which the acceptor is excited by FRET is proportional to the square of the scalar product of its transition dipole with the local electric field of the donor transition dipole. This is the origin of the k2 term in equation (8.3), given by: k2 ¼ ½ p^D p^A  3ð p^D^r DA Þð^r DA p^A Þ2 ¼ ðcos YT  3 cos YD cos YA Þ2 ð8:5Þ where p^D and p^A are the donor and acceptor transition dipole moment vectors, ^r DA is the vector between their centers, and the angles YT, YD, and YA are defined in Fig. 8.2A. k2 can take values between 0 and 4, as shown in Fig. 8.2B. If the value is not known, it becomes very hard to extract distances from FRET efficiencies. However, if at least one fluorophore is flexible, so that it experiences many orientations during the lifetime of the excited state, then k2 should average to 2/3. In the great majority of studies this assumption has been made. Where fluorescein is tethered to a phosphate group of RNA via a flexible linker, this is probably a good approximation. The negative charge is repelled by the backbone so that the fluorophore freely rotates in a cone, and the fluorescent anisotropy of fluorescein attached to DNA is typically low (usually
0.1). By contrast, the cyanine fluorophores interact strongly with DNA and RNA. We have shown that both Cy3 (Norman et al., 2000) and Cy5 (Iqbal

A D ΘD

f

ΘT

ΘA

A⬘

A

B

k 2= 0

k 2= 1

k 2= 4

Figure 8.2 Orientation of transition moments of cyanine fluorophores terminally attached to double-stranded DNA. (A) The orientation parameter k2. The transition dipole vectors for the coupled donor and acceptor fluorophore are indicated by the arrows, labeled D and A. Vector A0 is generated by the in-plane translation of vector A to share its origin with vector D. The definition of k2, given in Eq. (8.5), is based upon the angles shown. (B) If the fluorophores lie in parallel planes, the orientation parameter simplifies to k2 ¼ cos2 YT and varies between 0 and 1. The schematic shows the limiting cases, where the transition moments are parallel (k2 ¼ 1) and crossed (k2 ¼ 0). If the transition moments are colinear, k2 ¼ 4.

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et al., 2008a) when attached to 50 -termini via C3 linkers (as often generated when coupled as phosphoramidites during synthesis) stack upon the ends of double-helical DNA very much in the manner of an additional basepair (Fig. 8.3). The relative immobilization of these fluorophores under these conditions suggests that orientation could be an important factor in FRET efficiency, and because this pair is commonly used in single-molecule studies, this could have significant practical consequences. If Cy3 and Cy5 are terminally attached to a duplex DNA or RNA molecule, the NMR structures suggest that their planes would be approximately parallel and coaxial, but the angle between their transitions moments (those are approximately directed along the polymethyne linkers between the two indole rings; Iqbal et al., 2008a) will depend upon the length of the helix and its periodicity. k2 should be maximal (k2
1) when the transition dipoles are parallel, and minimal (k2
0) when they are perpendicular. Thus, it would be expected that EFRET should be modulated with twice the periodicity of the helix. This experiment was carried out for both DNA and RNA–DNA hybrid helices, using both ensemble and single-molecule methods (Iqbal et al., 2008b). For both DNA and RNA–DNA duplexes, the FRET efficiency falls with length, but with a pronounced modulation (Fig. 8.4). EFRET is modulated with twice the helical periodicity; for example, for RNA there are clear maxima at 11 and 17 bp, with minima at 15 and 21 bp. The DNA helix exhibits the same behavior, but with a phase shift indicative of a more tightly wound helix. This is exactly the behavior expected for orientation dependence of FRET. However, the modulation is clearly

Figure 8.3 Molecular graphics image showing the structure of Cy3 stacked onto the terminal basepair of a DNA duplex, as determined by NMR (Norman et al., 2000).

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1

0.8 RNA/DNA

E FRET

0.6

DNA

0.4

0.2

0

10

12

14

16 18 20 Duplex length/bp

22

24

Figure 8.4 Efficiency of energy transfer for Cy3 þ Cy5-labeled hybrid DNA and RNA/DNA duplexes as a function of duplex length (Iqbal et al., 2008b). EFRET was measured for each duplex species as phospholipid vesicle-encapsulated single molecules. The EFRET values are plotted for the DNA (squares) and DNA/RNA duplexes (circles) as a function of helix length, with the estimated errors. The lines show simulation of the data, using a model in which the major fraction of the fluorophores was stacked onto the helix undergoing lateral motion. For the DNA duplexes this was based on standard B geometry with a periodicity 10.5 bp/turn and a helical rise of ˚ /bp step; 31% of the fluorophore was allowed to be freely mobile (based on time3.6 A resolved analysis) with k2 ¼ 2/3, while the remaining fluorophore underwent lateral motion with a Gaussian half-width of 42 . For the DNA/RNA duplexes, the simulation was based on standard A geometry with a periodicity 12 bp/turn and a helical rise of ˚ /bp step; 12% of the fluorophore was allowed to be freely mobile (based on time3A resolved analysis) with k2 ¼ 2/3, while the remaining fluorophore underwent lateral motion with a Gaussian half-width of 42 . The fluorescent quantum yield for Cy3 was 0.30 attached to DNA and 0.35 attached to DNA/RNA. The refractive index was 1.33. The phase shift between the two helical forms is very clear. Single duplex molecules were studied trapped within phospholipid vesicles in 10 mM Tris–HCl (pH 8.0), 50 mM NaCl. Individual DNA or RNA/DNA duplex species were encapsulated in phospholipid vesicles comprising a 100:1 mixture of either L-a-phosphatidylcholine or 1,2dimyristoyl-sn-glycero-3-phosphocholine with 1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine-N-(cap biotinyl). The vesicles were attached to quartz slides coated with biotin-functionalized polyethylene glycol, via NeutrAvidin. Encapsulated molecules were excited at 532 nm by prism-based total internal reflection.

incomplete (EFRET does not become zero at the minima), suggesting that the orientation is subject to averaging by a combination of lateral mobility of the fluorophores on the end of the helix, and torsional mobility of the helices. The data were well simulated by a model based on the geometric

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properties of standard B- and A-form helices for DNA and RNA–DNA, and the positions of the fluorophores determined by NMR with a significant lateral averaging (Iqbal et al., 2008b), with or without a fraction of unstacked fluorophore as indicated by time-resolved studies (Iqbal et al., 2008b; Sanborn et al., 2007). This is not only a nice confirmation of the expected orientational dependence of FRET, but also a warning that such effects could significantly affect the interpretation of FRET data in terms of distance with a commonly used pair of fluorophores. Similar results have been obtained using more immobilized fluorophores (Bo¨rjesson et al., 2009; Lewis et al., 2005), but it is clear that the intrinsic immobility of the cyanine fluorophores is sufficient to result in significant variation of k2. Distances for ˚ if it is assumed that k2 ¼ 2/3, the duplex systems could be in error by 12 A where the largest discrepancy arises when the fluorophores are perpendicular (i.e., at the minima of EFRET). If the fluorophores are not constrained to lie in parallel planes, then k2 could reach a value of 4 in principle, so the potential error could be still greater. It is possible to imagine, for example, that states might be misassigned in a single-molecule time profile. The orientation dependence of FRET leads to something of a conundrum. If the fluorophores are mobile then k2 ¼ 2/3 may be a good approximation, and the distances between the fluorophores can be determined. But because of the flexibility, the position of the fluorophores relative to the RNA is not well known, so the interpretation of the distance in terms of the RNA structure is difficult. On the other hand, if the fluorophores are fixed so that their position on the RNA is known, this leads to a potential uncertainly with k2, so complicating the calculation of the distance. There are two ways to deal with this problem. One approach is to maximize the mobility—by selecting mobile fluorophores such as fluorescein and flexible linkers. In many cases, we are trying to distinguish between conformations such that the difference in donor–acceptor distance may be greater than the uncertainty in position. It may be possible to determine the average position of the mobile fluorophore, by reference to a less mobile fluorophore in a series of positions on a known structure (Norman et al., 2000), or using molecular dynamics calculations (Wozniak et al., 2008). Alternatively, if immobile fluorophores are chosen then the orientation dependence can potentially provide valuable angular information for structural modeling. It is likely that this will be exploited much more in the near future.

3. Choice of Fluorophores Small-molecule extrinsic fluorophores are generally used in structural and folding studies of RNA molecules (Fig. 8.5). There are a number of factors that will influence the choice of donor–acceptor fluorophores

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HO

O

(H3C)2N

O

O

N(CH3)2

CO2

CO2

R

R

Tetramethylrhodamine

Fluorescein

N

N n

R⬘

R

Cyanine N rib

O

N

H N

N H

H

S N

N H O

N rib

1,3-diaza-2-oxophenothiazine

Figure 8.5 The chemical structures of some fluorophores commonly used in FRET experiments. 1,3-Diaza-2-oxophenothiazine is a cytosine analog (Wilhelmsson et al., 2001), and is shown basepaired to guanine (gray).

for FRET studies in RNA. One is the distance range to be explored. Given the dimensions of natural macromolecules it is good to choose a fluorophore pair that allows measurements up to 70 A˚ or more, requiring an R0 ˚ or greater. The spectral overlap integral for Cy3–Cy5 is J (2/3) of 55 A (l) ¼ 7.2  10 13 M 1 cm 3 (Iqbal et al., 2008b) (Fig. 8.6), giving R0 ˚ ; thus EFRET falls (2/3) ¼ 60.1 A˚. For fluorescein–Cy3, R0 (2/3) ¼ 56 A from 0.8 to 0.2 (a conservative range for the measurement of FRET) over ˚ . This is a very useful size range for the study of the distance rDA ¼ 44–70 A RNA folding. The orientation of the fluorophores is clearly another important factor. This may depend on the intrinsic properties of the fluorophores and the manner of their attachment to the nucleic acid. As discussed above,

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3 ´ 105

0.6 Cy3 fluorescence

Cy5 absorption

2.5 ´ 105

0.4

2 ´ 105

0.3

1.5 ´ 105

0.2

1 ´ 105

0.1

0.5 ´ 104

0 500

550

600 650 Wavelength/nm

700

Cy5 absorption

Cy3 fluorescence

0.5

0 750

Figure 8.6 The fluorescence emission spectrum of Cy3 and the absorption spectrum of Cy5, showing the overlap between them.

fluorescein is quite mobile when it is attached to the 50 -termini of DNA or RNA. The cyanine fluorophores are predominantly stacked on the end, with a fraction that is unstacked at any moment (Iqbal et al., 2008b; Sanborn et al., 2007). This results in relatively complicated photophysical properties with multiple lifetimes. The dynamic and spectroscopic properties of tetramethylrhodamine are also quite complex (Neubauer et al., 2007; Va´mosi et al., 1996). If a fixed orientation of the fluorophores is required, then it may be advantageous to use fluorescent base analogs that are fixed in the helix by basepairing. The cytosine analog 1,3-diaza-2-oxophenothiazine is fluorescent and reasonably bright, but can basepair with guanine normally (Wilhelmsson et al., 2001) (Fig. 8.5). Wilhelmsson and colleagues (Bo¨rjesson et al., 2009) have recently synthesized two analogs 1,3-diaza-2oxophenoxazine (tCO) and 7-nitro-1,3-diaza-2-oxophenothiazine (tCnitro) that can act as a FRET donor–acceptor pair, with a calculated R0 (2/3) of ˚ . Although this is a relatively short Fo¨rster length, the base analogs can 27 A be placed quite close within the structure. It would be expected that energy transfer between tCO and tCnitro would be strongly affected by orientation, and this has been demonstrated for a series of DNA duplexes (Bo¨rjesson et al., 2009). These fluorophores have not been synthesized as ribonucleoside analogs to date, but this should be possible.

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Fluorophores may be attached to RNA in a variety of ways. If the RNA is chemically synthesized, then some of the fluorophores (including fluorescein and Cy3) may be coupled to the 50 -termini as phosphoramidites. Alternatively, fluorophores may be coupled to a 20 -aminoribose incorporated into the RNA at a chosen point. A greater variety of fluorophores may be conjugated to aminolinkers as N-hydroxysuccinimide esters, either at the termini or at the 5 position of dU (assuming that a 20 -deoxyribose substitution can be tolerated at that position). In principle, fluorophores could be attached via the backbone phosphate groups, as demonstrated in DNA (Ozaki and Mclaughlin, 1992). At the present time the photophysical properties of fluorophores that have been conjugated postsynthetically to nucleic acids have not been well characterized in general.

4. Construction of Fluorophore-Labeled RNA Species In this laboratory RNA species are synthesized using ribonucleotide phosphoramidites with 20 -tert-butyldimethylsilyl protection. Fluorophores may be coupled to the 50 -terminus as phosphoramidites, with an average coupling efficiency of 97%. Alternatively, we conjugate fluorophores as N-hydroxysuccinimide esters to primary amine groups that have been incorporated as terminal or internal amino-linkers. After deprotection, the RNA is desalted by gel filtration followed by ethanol precipitation. All RNA species are purified by gel electrophoresis in polyacrylamide gels (usually 20%) containing 7 M urea. Fluorescently labeled species are significantly retarded, so that bands may be excised and the oligonucleotides electroeluted into 8 M ammonium acetate, and recovered by ethanol precipitation. Following the gel purification, it is very important to introduce a further stage of purification by reversed-phase HPLC. We use a C18 column, eluted with a linear gradient of 100 mM ammonium acetate/ acetonitrile, with a flow rate of 1 ml/min. The donor–acceptor-labeled RNA is then constructed by hybridization. The required stoichiometric combinations of fluorophore-labeled and unlabeled strands are mixed together in 90 mM Tris–borate (pH 8.3), 25 mM NaCl for 10 min. They are then placed at 80  C, followed by slow cooling over several hours. The doubly labeled species are purified by electrophoresis in a polyacrylamide gel under nondenaturing conditions at 4  C at 150 V in 90 mM Tris–borate (pH 8.3), 25 mM NaCl with recirculation at >1 L/h. The fluorescent species are recovered by band excision and electroelution.

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5. Steady-State Measurements of FRET Steady-state fluorimetry has been extensively used to analyze folding in nucleic acids, including RNA. FRET efficiency can be measured from the quenching of the donor, or the enhanced emission from the acceptor. It can also be measured from the depolarization of the acceptor fluorescence. But of these the most sensitive method is to measure acceptor emission. For this we find Clegg’s normalized acceptor ratio (Clegg, 1992), the most straightforward means to extract EFRET, requiring a minimal number of samples. I have written a program in MATLAB to carry out this analysis. In this approach two emission spectra are recorded from a donor–acceptor labeled RNA sample, with excitation at two wavelengths (n1 and n2). In the general case (e.g., fluorescein–Cy3), the emission at a given wavelength of a sample excited primarily at the donor wavelength contains emission from the donor, emission from directly excited acceptor and emission from acceptor excited by energy transfer from the donor: Fðn1 n0 Þ / feD ðn0 ÞFA ðn1 ÞEFRET da þ eA ðn0 ÞFA ðn1 Þa þ eD ðn0 ÞFD ðn1 Þd½ð1  EFRET Þa þ ð1  aÞg 0

ð8:6Þ

0

¼ F ðn1 n Þ þ F ðn1 n Þ A

D

where d and a are the molar fraction of molecules labeled with donor and acceptor, respectively. Superscripts D and A refer to donor and acceptor, respectively. eD(n0 ) and eA(n0 ) are the molar absorption coefficients of donor and acceptor, respectively, and FD(n1) and FA(n1) are the fluorescent quantum yields of donor and acceptor, respectively. Thus, the spectrum contains components due to donor emission (FD(n1n0 ), i.e., the final term containing FD(n1)) and those due to acceptor emission (FA(n1n0 ), i.e., the first two terms containing FA(n1)). A spectrum of an RNA sample labeled only with donor is normalized to the donor peak and subtracted, leaving just the acceptor components, that is, FA(n1n0 ). The pure acceptor spectrum thus derived is then normalized to one from the same sample excited at a wavelength (n00 ) at which only the acceptor is excited, with emission at n2. This gives the normalized acceptor ratio:   F A ðn1 n0 Þ EFRET deD ðn0 Þ eA ðn0 Þ FA ðn1 Þ ðratioÞA ¼ A ð8:7Þ ¼ þ A 00 F ðn1 n00 Þ eA ðn00 Þ e ðn Þ FA ðn2 Þ EFRET is directly proportional to (ratio)A, and can be easily calculated since eD(n0 )/eA(n00 ) and eA(n0 )/eA(n00 ) are measured from absorption spectra, and FA(n1)/FA(n2) is unity when n1 ¼ n2. An analogous normalization procedure for the measurement of efficiency from donor deactivation has been presented (Clegg, 1992) and used successfully for the analysis of the global

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structure of a four-way DNA junction (Clegg et al., 1992). However, it is generally less sensitive than the acceptor ratio method. In this laboratory we use an SLM-Aminco 8100, equipped with Glan-Thompson polarizers. The electronics have been updated by the ISS Phoenix system. Measurements of FRET efficiency are performed under photon counting conditions, with the polarizers crossed at the magic angle (54.7 ) to remove polarization artifacts. Fluctuation of lamp intensity is corrected using a concentrated rhodamine B solution as a quantum counter. Ensemble steady-state measurements of FRET have been extensively used to study branched RNA molecules, including the three-way HS1HS7HS3 junction of the hammerhead ribozyme (Bassi et al., 1997; Penedo et al., 2004; Tuschl et al., 1994), the 2HS5HS3 (Lafontaine et al., 2001), and HS1HS5HS2 (Lafontaine et al., 2002) junctions of the VS ribozyme, a number of four-way 4H junctions (Walter et al., 1998a), including that of the hairpin ribozyme (Walter et al., 1998a,b) and its hinged equivalent lacking two arms (Walter et al., 1998c), and more complex junctions such as the 2HS12HS2 junction found in the HCV IRES (Melcher et al., 2003). The 4H junction derived from the IRES junction folds as an archetypical perfect four-way RNA junction (Fig. 8.7) (Melcher et al., 2003). The junction was constructed with four arms each of 12 bp, with fluorescein and Cy3 attached to the 50 -termini as phosphoramidites during synthesis. Six vectors were made, corresponding to the six possible end-to-end distances. In the presence of 10 mM Mg2þ ions, two of the vectors exhibited significantly more efficient energy transfer than the other four, consistent with a pronounced antiparallel structure based upon coaxial stacking of A on B and C on S arms (Fig. 8.7A). The structure of the junction is dependent on the concentration of divalent ions. The FRET efficiency of each vector changes with Mg2þ ion concentration (Fig. 8.7B); the data can be fitted to a two-state folding model: unfold EFRET ¼ EFRET þ DEFRET

KA ½Mg2þ n 1 þ KA ½Mg2þ n

ð8:8Þ

unfold is the FRET efficiency of the vector in the absence of Mg2þ where EFRET ions, DEFRET is the change in FRET efficiency with ion-induced folding, KA is the apparent association constant for Mg2þ ion binding, and n is the Hill coefficient. The data were fitted by nonlinear regression, giving ½Mg2þ 1=2 ¼ 150 mM and n ¼ 0.9, where ½Mg2þ 1=2 ¼ ð1=KA Þ1=n : Conformational changes in RNA structure can also be induced by protein binding, and FRET may be used to follow such a folding process. A good example of this is provided by the k-turn. The k-turn motif comprises a three-nucleotide bulge followed by trans sugar-Hoogsteen GA pairs (Klein et al., 2001), and introduces a very tight kink into the axis of helical RNA in the presence of Mg2þ ions (Goody et al., 2003).

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B

U A A U A U CGG GCC

A

GGG CCC

C

G C G C C G

S

A 0.5

E FRET

0.4

A

C

S

B

0.3 0.2 0.1 BA

CA

AS

BC

BS

CS

Vector

B

0.5 BC

SB

0.24

SB

BC

0.22

0.42

0.38

0.2 10−7

E FRET

E FRET

0.46

10−6

10−5

10−4

10−3

10−2

10−1

[Mg2+]/M

Figure 8.7 Steady-state FRET analysis of a 4H four-way junction derived from the 2HS12HS2 junction of the HCV IRES (Melcher et al., 2003). The central sequence of the junction is shown. The four arms are sequentially named A, B, C and S. Donor– acceptor-labeled vectors for FRET analysis are constructed with 50 -terminally attached fluorescein (donor) and Cy3 (acceptor) on selected helical arms, named according to those arms in that order. Thus, BA is labeled with donor on the end of arm B, and acceptor on A. (A) Histogram of the FRET efficiencies of the six end-to-end vectors.

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Folding into the kinked geometry requires the presence of metal ions, and this can be readily followed by FRET. This has been studied using a short RNA duplex, with a central k-turn motif and terminally attached fluorescein and Cy3 fluorophores (Goody et al., 2003). As the k-turn adopts its tightly kinked geometry, the fluorophores become closer and FRET efficiency increases. This can be combined with chemogenetic dissection of the RNA to determine interactions that are critical to the stability of the folded state (Liu and Lilley, 2007; Turner et al., 2005). K-turn folding may also be induced by the binding of L7Ae-related proteins (including the human 15.5 kDa protein)—a good example of induced fit (Turner et al., 2005). Titration of Archeoglobus L7Ae protein into the fluorescein–Cy3-labeled RNA solution brings about a marked increase in EFRET (Fig. 8.8A). Fitting to a stoichiometric binding model gives a dissociation constant of 65 pM, showing that the affinity is very high. However, to measure the affinity would require an undetectably low concentration of fluorescent RNA in order to be in equilibrium, so that this value is only an upper limit. FRET provided the means to measure this via the rates of association and dissociation. The association rate was measured by mixing 10 nM fluorescein–Cy3labeled RNA and 11 nM L7Ae protein in a stopped-flow mixer (Fig. 8.8B). The progress of the binding reaction was followed by the reduced fluorescence intensity of fluorescein at 515 nm, that is, as the fluorescein donor became increasingly quenched due to energy transfer to Cy3 as the RNA bound L7Ae and consequently folded into the kinked geometry. The data were fitted to two exponentials, with a faster rate of 0.59 s 1, corresponding to an association rate of kon ¼ 5.4  107 M 1 s 1. The dissociation rate was measured in the fluorimeter (this did not require fast kinetics methods), giving koff ¼ 0.002 s 1. From these rates we calculated a dissociation constant of KD ¼ 10 pM. It is possible to apply steady-state FRET measurements to larger constructs, the main limitation being the difficulty of preparing the RNA suitably labeled with donor and acceptor fluorophores. P-RNA (Smith et al., 2005) and ribosomes (Dorywalska et al., 2005) have been labeled by hybridization of fluorescent oligonucleotides. Clegg, Noller, and their collaborators have studied dynamic processes in the ribosome by steadystate FRET, using fluorophores attached to ribosomal proteins (Ermolenko et al., 2007a,b; Hickerson et al., 2005; Majumdar et al., 2005).

(B) FRET efficiency as a function of Mg2þ ion concentration for the SB and BC vectors. The data have been fitted to a two-state ion binding model. Fluorescence emission spectra were recorded at 4  C using an SLM-Aminco 8100 fluorimeter with modernized Phoenix electronics (ISS Inc., Champaign, IL, USA). Spectra were corrected for xenon lamp fluctuations and instrumental variations, and polarization artifacts were avoided by crossing excitation and emission polarizers at 54.7 .

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Figure 8.8 Steady-state FRET analysis of the formation of a kinked geometry by a k-turn-containing RNA on the binding of L7Ae protein (Turner and Lilley, 2008; Turner et al., 2005). The sequence of the RNA is shown, with 50 -terminal fluorescein on the top strand and 50 -terminal Cy3 on the bottom strand. (A) FRET efficiency as a function of L7Ae concentration (closed circles). These data have been fitted to a stoichiometric binding model. In a separate experiment, singly fluorescein-labeled RNA of the same sequence was titrated with L7Ae and the fluorescence anisotropy r measured. Note that this changes very little over the range of protein concentration, indicating that the mobility of the fluorescein is not significantly altered. (B) Reaction progress curve for the association of L7Ae with Kt-7 RNA using a stopped-flow rapid mixer. Intensity of fluorescein emission at 515 nm is plotted as a function of time after mixing RNA and protein to final concentrations of 10 and 11 nM, respectively. The decrease in fluorescein fluorescence reflects increased FRET efficiency resulting from the kinking of the RNA on protein binding. The data are fitted to two exponential functions (line).

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6. Time-Resolved Measurements of FRET The excited state of a fluorophore typically exhibits multiple lifetimes, so that the emission (I(t) at time t) decays with a rate:
 X t IðtÞ ¼ ð8:9Þ ai exp ti i where there are i lifetimes ti with amplitudes ai. In a FRET experiment, using an RNA species labeled with donor and acceptor, the lifetime of the donor excited state (tDA) will be shortened due to energy transfer to the acceptor, and this provides information on subpopulations of different conformational states within the ensemble: " # X ð1 X t t IDA ðtÞ ¼ drDA ð8:10Þ fn Pn ðRDA Þ ai exp  ti ti ðR0 =rDA Þ6 0 n i where there are n distributions (each of amplitude fn) of donor–acceptor distance distributions Pn(rDA). The distributions are generally assumed to be Gaussian. Most of the fluorophores used in FRET measurements have excitedstate lifetimes in the nanosecond range, although the cyanine fluorophores have shorter lifetimes. There are broadly two ways to measure fluorescent lifetimes, working in either the time or the frequency domain. In the former, the ensemble of molecules in solution is excited by a very short pulse of light (ideally a d-pulse), usually from a titanium:saphire laser. The intensity of the emission as a function of time following the excitation pulse is measured by time-correlated photon counting, and a decay curve generated. The donor fluorescence decay is deconvoluted from the instrument response, and then fitted to multiple exponentials from which the lifetime distributions can be calculated. The decay curve for Cy3 50 terminally attached to a 16 bp DNA molecule is shown in Fig. 8.9. The data were fitted to three exponential rates, corresponding to lifetimes of 390 ps, 1.04, and 1.91 ns (Iqbal et al., 2008b). Levitus and coworkers (Sanborn et al., 2007) have made a detailed study of the photophysics of Cy3 terminally attached to DNA. They conclude that the short lifetime corresponds to Cy3 that is unstacked from the end of the helix, and thus able to relax by cis–trans photoisomerization in the polymethyne linker between indole rings. Working in the frequency domain, the sample is excited by light whose intensity is sinusoidally modulated at high frequency (typically MHz). Light from a continuous wave laser (e.g., the 488 nm line from

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Figure 8.9 Time-resolved fluorescent lifetime analysis of Cy3 attached to doublestranded DNA (Iqbal et al., 2008b). Fluorescent decay curve for Cy3 attached to a 16 bp DNA duplex, showing the experimental data and the instrument response function (IRF), and the fit to three exponential functions (line). The decay curve was generated using time-correlated single-photon counting, after excitation by 200 fs pulses from a titanium:sapphire laser at 4.7 MHz.

an argon-ion laser to excite fluorescein) is modulated using an electrooptical modulator such as a Pockels cell. Because of the finite lifetime of the excited state, the emitted light is demodulated and phase shifted relative to the excitation. The data are plotted as the phase shift and modulation of the emitted light as a function of the modulation frequency. The donor–acceptor distance distributions were determined for k-turncontaining RNA as a function of Mg2þ concentration using this approach (Goody et al., 2003) (Fig. 8.10). The shift in both sets of curves with the increase in ionic concentration is apparent. The data required fitting to two Gaussian distributions of rDA distance to achieve an acceptable distribution of the residuals. These correspond to mean distances of ˚ , indicative of a tightly kinked geometry and 83 A˚ corresponding to 23 A an extended structure. At low of Mg2þ concentration, the extended structure was the major species, but the kinked geometry was dominant at high of Mg2þ concentration. Time-resolved FRET has been employed to study a number of RNA junctions, including the hammerhead ribozyme (Rueda et al., 2003), the hairpin ribozyme (Klostermeier and Millar, 2001; Walter et al., 1999), and the HCV IRES 2HS12HS2 junction (Melcher et al., 2003).

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Figure 8.10 Two populations of conformation of k-turn-containing RNA using timeresolved FRET in the frequency domain by phase fluorimetry (Goody et al., 2003). Donor–acceptor distance distributions were analyzed using the same fluorescein–Cy3 end-labeled species studied by steady-state FRET (Fig. 8.8). (A) Plots of phase shift and demodulation of the fluorescein emission as a function of the presence of 1 nM (closed circles) and 50 mM (open squares) Mg2þ ions as a function of the modulation frequency. The best fits to the experimental data using one or two Gaussian distributions of rDA lengths are shown by the broken and solid lines, respectively. (B) Two-Gaussian rDA distributions P(R) calculated from the fit to the data obtained in 1 nM Mg2þ ions. The major species (65%) corresponds to the longer rDA distance. The integrated areas under the curves total to 100%. (C) Two-Gaussian rDA distributions calculated from the fit to the data obtained in 50 mM Mg2þ ions. The major species (70%) now corresponds to the shorter rDA distance. These measurements were performed at 4  C using a

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7. Single-Molecule FRET The ability to detect single photons by devices of very high sensitivity and efficiency such as electron-multiplying charge-coupled device (EMCCD) chips and avalanche photodiode photodetectors has made it possible to carry out FRET experiments on single molecules (Ha et al., 1996, 1999a), and this has been applied to RNA in the last decade (Ha et al., 1999b; Zhuang et al., 2000). Very rapidly this approach has become probably the major way in which FRET is now applied to analysis of the folding and dynamics of RNA molecules. This is a very large subject in its own right, and a comprehensive coverage is not attempted here; there are other excellent reviews dedicated to this topic (Aleman et al., 2008; Ha, 2001; Joo et al., 2008; Pljevaljcic and Millar, 2008; Roy et al., 2008). Single-molecule FRET is performed on an inverted microscope, where the fluorescence is captured by the objective lens, and then split into donor and acceptor wavelengths using dichroic mirrors. There are two basic modes for doing this. One is to measure the burst of fluorescence from a free RNA molecule as it transiently diffuses through a confocal volume (Deniz et al., 1999). This uses essentially the same technology as fluorescence correlation spectroscopy. From the intensity at the donor and acceptor wavelengths, the efficiency of energy transfer can be measured, and a histogram constructed from many such events. However, because the molecule is typically observed for about a millisecond, no conformational dynamics are measured. This method has been used to study the conformation of the hairpin ribozyme (Pljevaljcic et al., 2004). Alternatively, the RNA can be immobilized on a slide, in which case single molecules can be studied for many seconds and conformational transitions observed. The time of observation is limited by photochemical effects on the fluorophores, but this can be extended with recently improved oxygen scavenging systems (Aitken et al., 2008; Rasnik et al., 2006). Again there are two choices. A wide-field approach can be used,

K2-Digital Phase Fluorimeter (ISS Inc., Champaign, IL, USA), with excitation at 488 nm from a vertically polarized, argon ion laser, intensity modulated at 39 frequencies between 4 and 300 MHz. Donor emission was measured using a 10 nm bandpass filter centered at 520 nm to exclude scattered incident light and acceptor fluorescence and a polarizer set at 54.7 to remove instrumental artifacts. Measurements were referenced to fluorescein in 10 mM NaOH, with a lifetime of 4.05 ns. Phase shift and modulation data were analyzed with the parameter estimation program CFS_LS ( Johnson and Faunt, 1992), according to the theory given in Melcher et al. (2003). Goodness of fit was evaluated by w2, the distribution of residuals and by a runs test. Confidence intervals (single standard deviation) were determined by the Bootstrap method (Efron and Tibshirani, 1993).

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where fluorescence from many single molecules is collected simultaneously using an EMCCD camera, or light from one molecule at a time can be analyzed, and the surface scanned using a nanopositioning stage. The latter allows higher time resolution, but as the data acquisition rates of cameras become faster this is less important. For the former, the excitation can be carried out using prism-based total internal reflection microscopy, where the surface-bound molecules are excited by the evanescent wave. This requires an objective lens with a high numerical aperture. Or epi-illumination can be used, where the excitation passes through the objective lens (Sase et al., 1995). There are a number of ways in which RNA can be tethered to the surface for observation. Most require that one strand of the RNA carries a biotin covalently attached to its 50 -terminus, which can be accomplished during synthesis. This may then be bound via a streptavidin molecule to biotinylated BSA coating the surface of the slide, or alternatively to PEG. A way of avoiding a direct connection with the surface while localizing the RNA on the slide is to use phospholipid vesicle encapsulation (Boukobza et al., 2001; Okumus et al., 2004). In this approach the molecule can diffuse freely within the confines of a 200 nm diameter vessel (a volume of 4  10 18 L). The data shown in Fig. 8.4 were collected in this manner. A relatively early application of single-molecule FRET was to a ribosomal three-way RNA junction (Ha et al., 1999). The adenine riboswitch is a more complex three-way junction, an HS2HS3HS8 junction (Mandal and Breaker, 2004). Structural transitions were observed by measurement of FRET between fluorophores tethered to 5-amino-allyluridine nucleotides incorporated into the terminal loops (Lemay et al., 2006). The RNA exhibits repeated transitions between two states of high (EFRET ¼ 0.9) and low (EFRET ¼ 0.25) FRET efficiency (Fig. 8.11). The high FRET state would be consistent with the loop–loop interaction observed in the crystal (Serganov et al., 2004). In the absence of added metal ions, the riboswitch remains largely in the open (low-FRET) state, but the folded (high-FRET) state becomes stabilized as the ionic concentration is raised. The dynamics were altered upon binding its adenine ligand. Singlemolecule FRET has also been applied to four-way RNA junctions (Hohng et al., 2004), the junction-containing hairpin ribozyme (Nahas et al., 2004; Okumus et al., 2004; Tan et al., 2003), and its minimal hinged form (Zhuang et al., 2002). The data for the full form of the hairpin ribozyme were interpreted in terms of a three-state folding process, in which the dynamics of the four-way junction juxtaposes the loops, which may then undergo a docking process. In a subsequent study, the conformational dynamics of the ribozyme were exploited to use FRET to follow cycles of cleavage and ligation in the active ribozyme (Nahas et al., 2004). This might be regarded as the first example of single-molecule ribozyme enzymology.

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Figure 8.11 Conformational transitions in the pbuE adenine riboswitch nucleotide binding domain (Mandal and Breaker, 2004) observed by single-molecule FRET (Lemay et al., 2006). The riboswitch was fluorescently labeled in the terminal loops, and tethered to the surface of a quartz slide via a biotin terminally attached to the third, open helix. Time records of FRET efficiency (50 ms integration time) as a function of elapsed time for single riboswitch molecules are shown at various Mg2þ ion concentrations. These data were recorded using a total internal reflection fluorescence microscope with 532 nm laser excitation and a back-illuminated electron-multiplying CCD camera (Andor iXon). The imaging buffer was 50 mM Tris–HCl (pH 8.1), 6% (w/w) glucose, 1% 2-mercaptoethanol, 0.1 mg/mL glucose oxidase, 0.02 mg/mL glucose catalase, and the indicated concentration of MgCl2. Measurements were performed at room temperature (22  C). Single-molecule FRET efficiency after background correction was approximated by IA/(IA þ ID), where IA and ID are the fluorescence intensities of the Cy5 acceptor and Cy3 donor, respectively.

Single-molecule studies have revealed that some branched RNA species exhibit surprisingly heterogeneous dynamics (Lemay et al., 2006; Tan et al., 2003; Zhuang et al., 2002), while others (such as a simple 4H RNA junction) do not. Rates of conformational transitions can vary a hundred

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fold from molecule to molecule; these properties are very persistent, with very infrequent interconversion of kinetic properties by a given molecule. Suspicion initially focused upon possible inhomogeneity of the surface to which the molecules were bound. However, this possibility is excluded because the same heterogeneity in hairpin ribozyme molecules was found when they were encapsulated in phospholipid vesicles (Okumus et al., 2004). So, it seems that the heterogeneity may be somehow built into the structure of more complex RNA molecules. This is not well understood at the present time. Single-molecule FRET has been applied to the folding of the group I intron ribozyme (Lee et al., 2007a; Russell et al., 2002; Zhuang et al., 2000), the group II intron ribozyme (Steiner et al., 2008), the VS ribozyme (Pereira et al., 2008), and the interaction of a tetraloop and its receptor (Hodak et al., 2005). It has been used to study folding of telomerase RNA (Stone et al., 2007) and dynamic processes in the ribosome during translation (Blanchard et al., 2004a,b; Cornish et al., 2008, 2009; Lee et al., 2007b; Wang et al., 2007). As the systems become more complex, the profiles of FRET efficiency with time can become more complicated, with many states interconverting. Hidden Markov modeling algorithms have been used to uncover the states within such profiles (McKinney et al., 2006). In a recent development, FRET has been used to study the dynamics of single DNA junctions while under the application of stretching force (Hohng et al., 2007); this experiment should be directly applicable to branched RNA species. Single-molecule FRET has become an extremely powerful way to study the dynamics of RNA in complex assemblies. The main limitation to this is now probably the ease with which fluorescent labels can be incorporated where needed.

ACKNOWLEDGMENTS I thank my collaborators and coworkers who have contributed to the studies discussed in this review, including Bob Clegg, Tim Wilson, Taekjip Ha, Carlos Penedo, Jo Ouellet, Sonya Melcher, Terry Goody, Ben Turner, David Norman, and Daniel Lafontaine. Cancer Research UK is acknowledged for financial support of the work of this laboratory.

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