Single-Molecule RNA Science - Xiaowei Zhuang - Harvard University

Annu. Rev. Biophys. Biomol. Struct. 0.0:${article.fPage}-${article.lPage}. Downloaded from arjournals.annualreviews.org 

by HARVARD COLLEGE on 04/05/05. For personal use only. 

10 Feb 2005 13:0 AR AR243-BB34-17.tex XMLPublish SM (2004/02/24) P1: JRX 

AR REVIEWS IN ADVANCE10.1146/annurev.biophys.34.040204.144641 

(Some corrections may occur before final publication online and in print) 

I N 

R E V I E W S 

A D V A N C E 

SINGLE-MOLECULE RNA SCIENCE 

Annu. Rev. Biophys. Biomol. Struct. 2005. 34:399–414 

doi: 10.1146/annurev.biophys.34.040204.144641 

Copyright c○ 2005 by Annual Reviews. All rights reserved 

First published online as a Review in Advance on March 3, 2005 

Xiaowei Zhuang 

Department of Chemistry and Chemical Biology and Department of Physics, Harvard 

University, Cambridge, Massachusetts 02138; email: zhuang@chemistry.harvard.edu 

KeyWords single-molecule experiments, RNA folding, RNA catalysis, 

fluorescence resonance energy transfer, optical tweezers 

■ Abstract The development of single-molecule detection and manipulation has 

allowed us to monitor the behavior of individual biological molecules and molecular 

complexes in real time. This approach significantly expands our capability to characterize 

complex dynamics of biological processes, allowing transient intermediate 

states and parallel kinetic pathways to be directly observed. Exploring this capability 

to elucidate complex dynamics, recent single-molecule experiments on RNA folding 

and catalysis have improved our understanding of the folding energy landscape of RNA 

and allowed us to better dissect complex RNA catalytic reactions, including translation 

by the ribosome. 

CONTENTS 

INTRODUCTION .....................................................399 

EXPERIMENTAL TECHNIQUES ........................................401 

Single-Molecule FRET ................................................401 

Optical Tweezers.....................................................403 

SINGLE-MOLECULE FRET STUDIES OF RNA FOLDING ..................403 

Folding of Large, Multidomain RNA Enzymes .............................403 

Folding of Small RNA Enzymes ........................................405 

Single-Molecule �-Value Analysis ......................................406 

SINGLE-MOLECULE FRET STUDIES OF RNA CATALYSIS .................407 

Catalysis by a Small RNA Enzyme ......................................407 

Translation by the Ribosome ...........................................407 

SINGLE-MOLECULE FORCE STUDIES OF RNA FOLDING 

AND UNFOLDING ...................................................408 

Folding and Unfolding of Simple Secondary or Tertiary Structures .............408 

Unfolding of a Large, Multidomain RNA Enzymes .........................409 

CONCLUSIONS AND FUTURE DIRECTIONS .............................409 

INTRODUCTION 

This is not the first time that we have witnessed RNA taking center stage. Some 

20 years ago, two independent, groundbreaking discoveries changed our view 

about this biomacromolecule: Found to function as catalysts, RNA molecules could 

1056-8700/05/0609-0399$20.00 399





400 ZHUANG 

no longer be viewed as merely messengers that pass information from the DNA 

to proteins (19, 28). Since then, a growing number of essential reactions in cells 

are have been found to be catalyzed by RNA enzymes. Two examples exist at the 

heart of the central dogma of biology: The ribosome, the giant ribonucleoprotein 

assembly responsible for protein synthesis in cells, uses its RNA components to 

perform major catalytic functions (12, 37, 38). The spliceosome also likely uses 

its RNA constituent to catalyze the splicing reaction of the precursor RNA (70, 

71), while the 100 or so protein splicing factors regulate the reaction. In addition 

to these essential catalytic functions, we now witness a second revolution in the 

world of RNA: its remarkable regulatory role. In the past decade or so, hundreds 

of small noncoding RNAs, microRNAs, and endogenous small-interfering RNAs 

were discovered that regulate gene expression either by inducing messenger RNA 

degradation or by suppressing translation in both prokaryotic and eukaryotic cells 

(3, 11, 17). Aside from these posttranscriptional regulations, gene-regulation by 

noncoding RNA at the transcription level has also been observed recently (17, 74). 

In keeping with this expanding list of fundamental biological roles, RNA also finds 

exciting applications in modern biotechnology and medicine, as gene regulatory 

tools, as therapeutic agents, and in drug discovery (9, 14, 60, 64). 

Although RNA science is attracting a tremendous amount of attention, singlemolecule 

studies of RNA are also gaining momentum. There are good reasons for 

this to happen. With a growing appreciation of the importance of RNA, a mechanistic 

understanding of how RNA molecules adopt specific structures that mediate 

their functions is becoming more desirable than ever. Research on this RNA folding 

problem suggests that the free energy landscape of RNA is often highly rugged (8, 

24, 41–43, 45, 46, 50, 52–54, 56, 66, 67, 69, 78–82). As a result, an RNA molecule 

can traverse a multitude of kinetic paths and intermediate conformational states 

before attaining its native structure. For example, oligonucleotide hybridization, 

hydroxyl radical footprinting, circular dichroism, and UV absorption assays have 

led to the discovery that RNA folding occurs through a series of intermediate 

states (46, 56, 69, 79, 80). RNA enzymes were also seen to fold along multiple 

pathways by native gel electrophoresis and enzymatic activity assays (41–43, 

52). However, these methods that measure the average property of an ensemble of 

molecules can only probe accumulative intermediate states and significantly populated 

pathways; they cannot detect nonaccumulative, transient intermediate states. 

The existence of a large number of folding pathways will also make the characterization 

of each individual pathway difficult by ensemble assays. In these cases, 

single-molecule approaches come to the rescue owing to their intrinsic capability 

to detect nonaccumulative transient intermediate states and heterogeneity in the 

system. 

Similarly, the reactions catalyzed by RNA enzymes, especially those by large 

ribonucleoprotein complexes such as the ribosome, tend to be complex, involving 

many reaction steps and reaction intermediates. These steps are typically not 

synchronized for an ensemble of molecules, making the reactions difficult to dissect. 

Although stalling the reaction complexes at specific intermediate states has





SINGLE-MOLECULE RNA SCIENCE 401 

provided us with a lot of molecular insights into these reactions, this kind of 

manipulation is not necessarily possible for every intermediate state. Monitoring 

the reaction of a single molecule or molecular complex naturally overcomes this 

synchronization problem, offering us an opportunity to resolve each microscopic 

step of a complex reaction. 

The power of single-molecule techniques to elucidate complex dynamics does 

not come without sacrifices. Because of the limited amount of signal that a single 

molecule can produce, the time and spatial resolutions of single-molecule experiments 

are relatively limited compared with ensemble experiments. Luckily, RNA 

folding and catalytic reactions are not so fast, typically occurring on the timescales 

of milliseconds to minutes. In addition, these reactions often involve large conformational 

changes. These properties allow current, state-of-the-art single-molecule 

experiments to explore the RNA world. Indeed, these experiments have already 

made significant contributions to our understanding of the structural biology and 

enzymology of RNA. In this article, I review some of the recent developments in 

RNA science made possible by single-molecule techniques, focusing on those by 

fluorescence resonance energy transfer (FRET) and optical tweezers. 

EXPERIMENTAL TECHNIQUES 

Single-Molecule FRET 

FRET is a powerful assay to measure the intra- and intermolecular motions of 

biomolecules in real time (57, 63, 75). In this assay, a pair of fluorescent donor and 

acceptor molecules is attached to the host biomolecule(s) of interest. The dipoledipole 

interaction between the donor and acceptor causes energy transfer between 

them, leading to a decrease in the donor emission and an increase in the acceptor 

emission. The energy transfer efficiency E is given by E = 1/(1 + (R/R0) 6 ), 

where R is the distance between the donor and the acceptor. The Förster radius R0 

is typically 3 to 8 nm, making FRET sensitive to changes of a few nanometers in 

the donor-acceptor distance, which can reflect either the conformation changes of 

the host molecule or relative motions between two molecules. The dependence of 

energy transfer efficiency on the orientation of the donor and acceptor fluorophores, 

however, introduces uncertainty into FRET-based distance determinations. This 

problem can be mitigated if the dyes are attached to the host molecule with flexible 

linkers to ensure their free rotation. As this condition is difficult to satisfy rigorously 

in practice, researchers should be cautious when translating FRET efficiencies into 

absolute distances quantitatively. 

The development of single-molecule fluorescence spectroscopy (33, 36, 77) 

has allowed FRET to be measured at the single-molecule level (21, 55). Here, 

I briefly discuss a few aspects of single-molecule detection that are critical for 

unveiling the dynamic information of biomolecules. Like all single-molecule 

fluorescence techniques, efficient reduction of the background signal is critical





402 ZHUANG 

for single-molecule FRET. This is often achieved by confocal detection or by 

excitation with an evanescent wave generated by total internal reflection. The total 

internal reflection microscope allows a whole field of molecules, typically several 

hundred of them, to be detected simultaneously. Such a parallel detection 

greatly facilitates the building of statistics, especially for irreversible processes. 

However, wide-field detection requires the use of cameras, which limit the data 

acquisition rate to 1 kilocycle per second or slower using state-of-the-art charge 

couple device (CCD) cameras with on-chip amplification. Higher data acquisition 

rates, up to tens of megacycles per second, can be achieved with confocoal 

microscopes using sensitive point detectors, such as avalanche photodiodes or 

photomultipliers. 

Besides physical instrumentation, special manipulation of biomolecules is often 

required for real-time characterizations. With a confocal microscope, it is possible 

to detect freely diffusing molecules in solution (13, 36). Although this detection 

geometry avoids any potentially adverse effect arising from the confinement of 

molecules, the range of timescales accessible by this method is severely limited 

by the dwell time of a molecule in the confocal detection volume, typically in the 

submilliseconds governed by diffusion. Therefore, to expand the dynamic range 

of the single-molecule experiments, various strategies were developed to confine 

the biomolecules of interest within the detection volume of a microscope. 

The two most commonly used strategies are confinement by matrices, such as 

agarose or polyacrylamide gels (32), and immobilization to surfaces with specific 

interactions, such as via the biotin-streptavidin bridge (22). The latter method is 

compatible with rapid buffer exchange and therefore is better suited for characterizing 

reactions. Although surface immobilization may perturb the biomolecules, 

this perturbation has been rather insignificant for most of the RNA systems studied 

to date, at neutral or basic pH values. For systems that involve proteins, which 

tend to interact strongly with surfaces through hydrophobic interactions, surface 

passivation is often required to retain the integrity of the system. Polyethylene 

glyco (PEG) is a commonly used passivating reagent (34). Another method is to 

entrap the biomolecule in a lipid vesicle attached to a surface (48). This technique 

potentially offers a close-to-physiological environment for the biomolecule; 

however, a method for rapidly changing the buffer in the vesicle needs to be 

developed before this method can be broadly used for characterizing biological 

processes. 

With the molecules confined in the detection volume, dynamics can be probed 

at the single-molecule level over a wide range of timescales, ranging from millisecond 

to minutes or even hours. The lower limit is imposed by the photon 

emission rate from a single fluorophore under conditions in which photoblinking 

or photobleaching is not severe enough to render the experiment impractical. 

The recent development of CCD cameras with on-chip amplification allows this 

limit to be achieved even in the wide-field-detection geometry. This limit can be 

further improved by using fluorescence correlation spectroscopy if the process 

under investigation is reversible and if the forward and backward reaction occurs






spontaneously (25, 27, 76). In this case, the time resolution is limited by the response 

time of the detectors, which is of the order of tens of nanoseconds for 

avalanche photodiodes or photomultipliers. 

Single-molecule FRET has been used to study a variety of biological systems. 

In this review, I only cover its application to RNA systems (4–7, 22, 35, 47, 51, 

54, 67, 78, 81, 82) but refer interested readers to a number of recent review articles 

for other applications (20, 75, 83). 

Optical Tweezers 

Optical tweezers rely on a tightly focused laser beam(s) to trap a particle in three 

dimensions and, through redirection of the beam, manipulate the particles (1). 

Focused light exerts two forces on the particle, the gradient force and the scattering 

force. When the refractive index of the particle is larger than that of the surrounding 

medium, the gradient force draws the particle toward the focus of the beam, where 

the light field is the strongest. The scattering force arises from the radiation pressure 

exerted on the particle by photons that are either absorbed or scattered. When 

balanced, these two forces hold the particle just slightly downstream of the light 

focus. 

Optical tweezers allow the manipulation and detection of single biomolecules. 

As biological molecules are often too small to be manipulated directly by optical 

tweezers, a micron-sized dielectric sphere is often attached to the molecule of 

interest to serve as a handle. This allows researchers to manipulate the position of 

the attached biomolecule and to exert a well-defined force. Both parameters can 

be determined with exquisite accuracy: The position of the microsphere can be 

measured to within 0.1 nm, and the force to 0.1 pN (39, 58). Forces well above 

100 pN can be achieved with a dual-beam optical tweezers (10). 

Since the invention of optical tweezers, this technique has been adopted to 

study a variety of biological systems. This review focuses on the studies on RNA 

systems (23, 30, 31, 40). Other single-molecule force spectroscopy techniques, 

such as those using atomic force microscopes and magnetic tweezers, have also 

provided a wealth of information on biological molecules (61, 62, 83). These 

single-molecule techniques are fully expected to make significant contributions to 

our understanding of RNA systems as well. 

SINGLE-MOLECULE FRET STUDIES OF RNA FOLDING 

Folding of Large, Multidomain RNA Enzymes 

The capability of single-molecule FRET to follow conformational changes of 

RNA was clear early on. In one of the first applications of single-molecule FRET 

to biological systems, Ha et al. (22) showed that the ligand-induced conformational 

changes of an RNA three-way junction could be monitored in real time at 

the single-molecule level. Zhuang et al. (81) performed the first single-molecule





404 ZHUANG 

RNA folding study by using a large, multidomain RNA enzyme, the Tetrahymena 

ribozyme (Figure 1a, see color insert), as a model system. The power of singlemolecule 

FRET was already apparent in this early work. A repertoire of 

dynamic properties that are difficult to analyze with ensemble experiments, including 

nonaccumulative folding intermediate states, parallel folding pathways, 

and equilibrium conformational fluctuations, were directly observed in singlemolecule 

FRET trajectories of the Tetrahymena ribozyme (81). 

FRET trajectories of individual ribozyme molecules directly reveal folding 

intermediates (Figure 1b): The addition of Mg 2+ to trigger folding causes the 

FRET signal to increase from the unfolded state value of 0.1 to an intermediate 

value (0.3) before attaining the native value (0.9). Statistical analysis of the 

folding times of each molecule indicated that the Tetrahymena ribozyme folds 

along at least three distinct pathways, with folding rate constant being 1 s, 1 min, 

and 0.01 hr, respectively. Kinetic traps exist on these pathways. While the two 

slower pathways have been observed previously in ensemble measurements, the 

most rapid pathway was first observed in this single-molecule FRET experiment 

(81). 

In a follow-up study designed to gain a greater understanding of the various 

pathways (54), folding of the Tetrahymena ribozyme was started from distinct regions 

on the energy landscape by varying the monovalent salt concentration of the 

prefolding solution. Different monovalent salt concentrations led to dramatically 

different folding behaviors, each with a distinct folding rate constant and traversing 

a different set of intermediate states. These results were interpreted as the 

molecules folding along several discrete channels separated by large energy barriers. 

Site-specific mutation analysis suggests that these different folding behaviors 

are caused by the different secondary structures formed under various salt concentrations 

(54), consistent with previous ensemble experiments (43, 44). Recent 

results from a hydroxyl radical footprinting assay suggest that high concentrations 

of monovalent salt induce an extensive amount of tertiary structures in addition 

to secondary structure changes (66). These preformed tertiary structures may also 

be partly responsible for the observed changes in the folding kinetics induced by 

preincubation with monovalent salt. 

More recently, FRET has been applied to another large RNA molecule, the 

catalytic domain of Bacillus subtilis RNase P RNA (78). Ensemble characterizations 

of this enzyme indicate the existence of three conformational states: the 

unfolded state, the intermediate state, and the native state. However, analysis of 

the molecules at equilibrium using single-molecule FRET suggest that at least 

seven different conformations exist at intermediate Mg 2+ concentrations, as distinguished 

by their corresponding FRET levels and connectivity between different 

states. The equilibrium structural dynamics of individual RNase P molecules also 

shows a high level of heterogeneity. 

These single-molecule results indicate that large RNA molecules fold across 

a highly rugged energy landscape, along a multitude of folding pathways, and 

through many intermediate folding states. These findings not only corroborate






previous and concurrent ensemble characterizations using various footing-printing 

methods, enzymatic activity assays, gel-mobility measurements, and optical scattering 

or spectroscopic techniques (8, 24, 41–43, 45, 46, 50, 52, 53, 56, 66, 69, 

79, 80), but also reveal new folding intermediates and folding pathways difficult 

to access by ensemble assays, thereby significantly improving our understanding 

of RNA folding. 

Folding of Small RNA Enzymes 

To date, single-molecule studies of small RNA enzymes have concentrated on one 

type of ribozyme, the hairpin ribozyme (Figure 2a, see color insert), although other 

systems are beginning to be explored. The hairpin ribozyme has been suggested 

to assume either an extended, undocked conformation or a docked conformation 

in which the two loop domains, A and B, make tertiary contacts (73). Two forms 

of hairpin ribozymes have been studied at the single-molecule level: the two-way 

junction ribozyme, a minimal hairpin ribozyme with high enzymatic activity, and 

the wild-type ribozyme, which has a four-way junction connecting helix 2 and 

helix 3 (15, 73) (Figure 2a). The first single-molecule experiment was performed 

on the two-way junction ribozyme (82). Strikingly, the FRET time traces of single 

molecules show highly heterogeneous undocking kinetics with several different 

rate constants (Figure 2b,c). Furthermore, the molecules retain their particular 

undocking rates through many docking-undocking cycles, and conversion between 

different undocking behaviors requires at least several hours (Figure 2b). These 

results suggest the existence of rather large numbers of conformational states that 

are all relatively stable under functional conditions, which is remarkable for such 

a small RNA enzyme. 

The heterogeneous undocking kinetics appear to be ubiquitous to all hairpin 

ribozymes. For the two-way junction ribozyme, both the wild type and variants with 

significantly different docking and undocking rate constants show heterogeneous 

undocking behavior (7, 51, 82). In addition, heterogeneous docking kinetics are 

seen in a variant with a modification at the domain junction (51). A study on 

the four-way junction hairpin ribozyme, a construct that more closely resembles 

the natural form of the ribozyme, also shows pronounced heterogeneity in both 

docking and undocking kinetics (67). 

The comparison between the single-molecule studies of the two- and fourway 

junction ribozymes reveals an interesting phenomenon: The latter form folds 

(docks) two to three orders of magnitude faster (67). Evidence suggests that the 

dramatic enhancement in the folding rate is a result of the intrinsic structural 

dynamics of the four-way junction: The ribozyme appears to fold into its native 

form via an intermediate state in which the four-way junction juxtaposes the loop 

domains, A and B, into proximity but without substantial loop-loop interactions 

(67). This result is further supported by a more recent single-molecule study of 

hairpin ribozymes freely diffusing in solution (47). There, even when the two loops 

are replaced with perfectly base-paired helices, these helices are still juxtaposed





406 ZHUANG 

into proximity by the four-way junction, whereas both two-way and three-way 

junctions lack this capability. Undocking kinetics does not seem to be accelerated 

by the junction dynamics. 

These results suggest that even small RNA enzymes can have highly complex 

structural dynamics, featuring a rugged energy landscape in the conformational 

space. Unlike in the case of large, multidomain RNA molecules, the level of complexity 

in the structural dynamics of small ribozymes had not been observed previously 

in ensemble experiments. These single-molecule studies make us wonder 

whether rugged energy landscapes and complex structural dynamics might very 

well be general properties of all RNAs, not just reserved for the large, complex 

RNA enzymes. 

Single-Molecule �-Value Analysis 

Characterization of the transition state ensemble is critical for a mechanistic understanding 

of the macromolecule folding problem. While transition-state analysis 

has been performed routinely for protein folding, its implementation in RNA folding 

has been challenging owing to the rugged energy landscape of RNA, which 

often leads to the coexistence of distinct structural transitions. The power of singlemolecule 

measurements to isolate individual transitions makes them ideally suited 

for transition-state analysis of RNA folding, A single-molecule �-value analysis 

based on FRET was developed to characterize the degree to which specific intramolecular 

interactions are formed in the transition states of RNA folding (4, 

7), following the �-value analysis introduced by Fersht (16) for characterizing 

protein folding. 

Using FRET to monitor the folding and unfolding transitions of the hairpin ribozyme, 

in conjunction with metal-ion titrations and tertiary-contact-destabilizing 

mutations, Bokinsky et al. (7) showed that the transition state is compact with the 

Mg 2+ -mediated interactions between the two domains of the hairpin ribozyme 

formed at a strength similar to that in the folded state, whereas the native tertiary 

contacts are not substantially formed. The conformational entropy loss and the 

breaking of secondary structures may be the primary mechanisms limiting the 

folding rate of the ribozyme. The folding transition states of some proteins also 

feature native-like topology without substantial native tertiary contacts (29). 

This single-molecule �-value analysis is not only applicable to an elementary 

folding reaction of a small RNA enzyme, it can also be used to characterize the 

transition state of a local folding step of a large RNA enzyme (4, 81). After extensive 

effort to disrupt every single tertiary contact between the P1 duplex and catalytic 

core of the Tetrahymena ribozyme, Bartley et al. (4) have shown that not one of 

these tertiary contacts is formed in the transition state of P1-docking, which is 

believed to be the last folding step of the ribozyme. Their results suggest that P1docking 

is rate-limited by kinetic traps. This work demonstrated the potential of 

this single-molecule �-value analysis to characterize each individual folding step 

of a complex, multistep, multipath folding reaction.






SINGLE-MOLECULE FRET STUDIES OF RNA CATALYSIS 

Single-molecule methods are particularly well suited for detecting transient intermediate 

states and thereby resolving the microscopic steps of a complex reaction. 

This is demonstrated in the following two examples, which are at the two extremes 

of the ribozyme world, a small RNA enzyme and the ribosome. 

Catalysis by a Small RNA Enzyme 

Following the reaction of single hairpin ribozyme molecules using FRET, Zhuang 

et al. (82) have shown that this ribozyme catalyzes the cleavage reaction of its 

substrate in several distinct steps (Figure 2c): (a) substrate binding; (b) folding 

of the ribozyme-substrate complex into a catalytically active state (docking); 

(c) cleavage of the substrate; (d) the unfolding (undocking) of the ribozyme-product 

complex; and (e) release of the products. These results confirmed a previously 

proposed reaction pathway based on ensemble measurements (15, 72, 73). The 

determination of the rate constants of each step indicates that the overall cleavage 

reaction of the two-way junction ribozyme is primarily rate-limited by the two 

structural transition steps, (b) and (d), and by the internal equilibrium constants 

of the reversible cleavage step, (c) (82). The heterogeneous undocking kinetics 

mentioned above result in heterogeneous reaction kinetics. 

The rate-limiting mechanism of the four-way junction ribozyme is different. 

Cleavage appears to dramatically accelerate undocking in the case of the four-way 

junction ribozyme. As a result the undocking rate is faster than the ligation rate, 

and the overall cleavage reaction rate is primarily limited by the internal cleavage 

reaction (35). In contrast, the undocking rate constant of the two-way junction 

ribozyme bound to its natural cleavage products is much slower than the ligation 

rate constant (51; G. Bokinsky, N. Walter & X. Zhuang, unpublished data). This 

discrepancy between the two-way and four-way ribozymes indicates an interesting 

allostery effect that the junction has on the docked structure of the loop domains. 

Translation by the Ribosome 

Early works by Hochstrasser and coworkers (26, 65) have shown that surfaceimmobilized 

ribosome complexes retain their peptidyl transferase activity and can 

be detected at the single-complex level. These experiments gave researchers hope 

that single-molecule experiments might someday provide exciting new insights 

into the molecular mechanisms of protein synthesis in cells. 

This promise has been realized in a recent, tour de force experiment by 

Blanchard et al. (5, 6). To set the stage for new discoveries, these authors first 

demonstrated that surface-immobilized ribosomes are highly active in tRNA 

accommodation and translocation and in peptide-bond formation (6). This is nontrivial 

considering the complexity of this vast ribonucleoprotein. Indeed specific 

surface immobilization and extensive surface passivation are necessary.





408 ZHUANG 

Next, using single-molecule FRET to follow the accommodation of the tRNAelongation 

factor Tu-GTP ternary complex in real time, Blanchard et al. (5) resolved 

a multistep movement of the tRNA into the ribosome and identified the 

intermediate states in the tRNA-delivery pathway using antibiotics and nonhydrolyzable 

GTP analogs. Previously unknown intermediate states were discovered 

both in the initial codon-recognition steps and in the kinetic proofreading steps 

following GTP hydrolysis (Figure 3, see color insert). These results, complementing 

previous kinetic characterizations of translation (18, 49), have provided new 

insights into the structural basis for the initial cognate tRNA selection, kinetic 

proofreading, and thus for the fidelity of translation by the ribosome. Interesting 

dynamic tRNA fluctuations between the classical state and hybrid states have also 

been observed (6). The translocation of tRNAs from the classical to the hybrid 

state after peptide-bond formation has been suggested before; however, it is surprising 

that tRNA fluctuates spontaneously between these two states both before 

and after peptide-bond formation. This observation again highlights the capability 

of single-molecule experiments to resolve dynamic behaviors. 

SINGLE-MOLECULE FORCE STUDIES OF RNA 

FOLDING AND UNFOLDING 

Folding and Unfolding of Simple Secondary 

or Tertiary Structures 

The use of mechanical forces to induce folding and unfolding provides a few unique 

advantages in characterizing macromolecule folding: (a)Itallows the folding and 

unfolding reaction to occur along a well-defined reaction coordinate, the molecular 

end-to-end distance; and (b) the folding free energy can be readily measured (68). 

Liphardt et al. (31) first characterized unfolding and refolding of small RNA 

molecules induced by mechanical forces using optical tweezers. They have shown 

that, with a slow-enough force-loading rate, the folding and unfolding of secondary 

structures, such as an RNA hairpin or a three-helix junction, can occur in 

equilibrium. In contrast, for a molecule with tertiary contacts, such as the P5abc 

domain of the Tetrahymena ribozyme in the presence of Mg2+ , folding and unfolding 

are nonequilibrium processes even at the slowest accessible force-loading rate. 

Interestingly, the transition state for folding for secondary structure formation is 

dramatically different from that for tertiary structure formation. In the former case, 

the transition state is relatively far from the folded state on the reaction coordinate, 

i.e., the molecular end-to-end distance, indicating a relatively soft transition. 

In contrast the transition state for tertiary structure folding is close to the folded 

state, suggesting that tertiary structures are much more brittle than secondary 

structures. 

As promised, the free energy difference between the folded and unfolded 

states is readily calculated from the force-extension curves of molecules being 

folded or unfolded at equilibrium, and compare well with theoretical predictions.






Remarkably, the folding free energy can even be derived from force-extension 

curves of nonequilibrium (un)folding processes using a recently developed nonequilibrium 

statistical mechanics theorem, the Jarzynski’s equality (30). 

Unfolding of a Large, Multidomain RNA Enzymes 

More recently, Onoa et al. (40) extended mechanical unfolding to a large RNA 

enzyme, the Tetrahymena ribozyme (Figure 4a, see color insert). The forceextension 

curves of individual RNA molecules show several discrete unfolding 

steps (Figure 4b), indicating the existence of multiple unfolding intermediate states. 

The molecular interactions disrupted at each of these unfolding steps have been 

identified by studying progressively larger pieces of the Tetrahymena ribozyme, 

determining the number of nucleotides released at each step, and using mutations 

or antisense oligonucleotides to perturb specific interactions in the RNA. Interestingly, 

most of these unfolding barriers are imposed by tertiary interactions, even 

though the overall thermodynamic stability of the molecule largely arises from the 

secondary structures, namely the base-paired helices. Unlike thermal or solution 

unfolding, in which tertiary structures are dissolved before secondary structures, 

the partially unfolded intermediates observed in this mechanical unfolding experiment 

contain both secondary structures and tertiary contacts. 

Mechanical unfolding experiments have been used to characterize the secondary 

structure formation of RNA as large as the 16S ribosome RNA. Even in this case, 

well-defined unfolding intermediates are observed (23). 

CONCLUSIONS AND FUTURE DIRECTIONS 

A distinct advantage of single-molecule techniques is their capability to detect 

transient, nonaccumulative states and heterogeneous behavior. Such capabilities 

make these techniques well suited to investigate the structural dynamics and catalytic 

reactions of RNA molecules. Indeed, single-molecule experiments have 

already yielded important insights into RNA folding by detecting new folding intermediate 

states and pathways, and by revealing a daunting level of complexity 

in the conformational dynamics of both large and small RNA. Single-molecule 

experiments have also allowed us to better dissect the complex catalytic reactions 

of RNA enzymes by resolving individual reaction steps and discovering 

new reaction intermediates that are critical for understanding the reaction 

mechanisms. 

What we have witnessed in the past few years is likely just the tip of the iceberg. 

Future single-molecule studies will certainly make more significant contributions 

to RNA science. As a growing number of essential reactions in cells have now 

been catalyzed by ribonucleoprotein enzymes, a bright future direction for singlemolecule 

studies would be to characterize the folding, assembly, and enzymatic 

reactions of these important cellular ribonucleoproteins. The recent work on the





410 ZHUANG 

delivery of tRNA to ribosome gives us confidence that these complex ribonucleoproteins 

are amendable to single-molecule studies. 

Another exciting direction is to apply these single-molecule studies to live cells. 

After all, the cellular environment is different from that in solution. A complete 

understanding of RNA folding, catalysis, and regulation requires investigation in 

live cells. Recent studies show that the trafficking of individual messenger RNAs 

and viral RNA genes can be monitored in living cells (2, 59). Although these studies 

have provided important insights into the trafficking properties of RNA-protein 

complexes, the investigations relied on the attachment of many fluorophores to each 

complex to compete with the cell autofluorescence background, a strategy that is 

not necessarily applicable to other RNA studies. Information that can be extracted 

from a single fluorophore in living cells is rather limited with the current stateof-the-art 

technology. However, intensive effort is being invested into developing 

brighter fluorescent reporters for single-molecule studies in live cells. Indeed, 

several research centers have recently been elected to tackle this problem. I am 

optimistic that these efforts will make it possible to investigate RNA folding and 

catalysis and the regulatory roles of RNA at the single-molecule level in living 

cells. 

ACKNOWLEDGMENTS 

This work is supported in part by the ONR, NSF, and a Packard Science and 

Engineering Fellowship. I thank S.C. Blanchard and S. Dumont for providing 

Figures 3 and 4, respectively. 

The Annual Review of Biophysics and Biomolecular Structure is online at 

http://biophys.annualreviews.org 

LITERATURE CITED 

1. Ashkin A, Dziedzic JM, Bjorkholm JE, 

Chu S. 1986. Observation of a single-beam 

gradient force optical trap for dielectric particles. 

Opt. Lett. 11:288–90 

2. Babcock HP, Chen C, Zhuang X. 2004. 

Using single-particle tracking to study nuclear 

trafficking of viral genes. Biophys. J. 

87:2749–58 

3. Bartel DP. 2004. MicroRNAs: genomics, 

biogenesis, mechanism, and function. Cell 

116:281–97 

4. Bartley LE, Zhuang X, Das R, Chu S, Herschlag 

D. 2003. Exploration of the transition 

state for tertiary structure formation 

between RNA helix and a large structured 

RNA. J. Mol. Biol. 328:1011–26 

5. Blanchard SC, Gonzalez RLJ, Kim HD, 

Chu S, Puglisi JD. 2004. tRNA selection 

and kinetic proofreading in translation. Nat. 

Struct. Mol. Biol. 11:1008–14 

6. Blanchard SC, Kim HD, Gonzalez RLJ, 

Puglisi JD, Chu S. 2004. tRNA dynamics 

on the ribozome during translation. 

Proc. Natl. Acad. Sci. USA 101:12893– 

98 

7. Bokinsky G, Rueda D, Misra VK, Gordus 

A, Rhodes MM, et al. 2003. Singlemolecule 

transition-state analysis of RNA 

folding. Proc. Natl. Acad. Sci. USA 100: 

9302–7 

8. Buchmueller KL, Webb AE, Richardson 

DA, Weeks KM. 2000. A collapsed





non-native RNA folding state. Nat. Struct. 

Biol. 7:362–66 

9. Burgstaller P, Jenne A, Blind M. 2002. Aptamers 

and aptazymes: accelerating small 

molecule drug discovery. Curr. Opin. Drug 

Discov. Dev. 5:690–700 

10. Bustamante C, Bryant Z, Smith SB. 2003. 

Ten years of tension: single-molecule DNA 

mechanics. Nature 421:423–27 

11. Carrington JC, Ambros V. 2003. Role of 

microRNAs in plant and animal development. 

Science 301:336–38 

12. Carter AP, Clemons WM, Brodersen DE, 

Morgan-Warren RJ, Wimberly BT, Ramakrishnan 

V. 2000. Functional insights 

from the structure of the 30S ribosomal 

subunit and its interactions with antibiotics. 

Nature 407:340–48 

13. Deniz AA, Dahan M, Grunwell JR, Ha 

T, Faulhaber AE, et al. 1999. Singlepair 

fluorescence resonance energy transfer 

on freely diffusing molecules: observation 

of Forster distance dependence and 

subpopulations. Proc. Natl. Acad. Sci. USA 

96:3670–75 

14. Famulok M, Verma S. 2002. In vivoapplied 

functional RNAs as tools in proteomics 

and genomics research. Trends 

Biotechnol. 20:462–66 

15. Fedor MJ. 2000. Structure and function of 

the hairpin ribozyme. J. Mol. Biol. 297: 

269–91 

16. Fersht A. 1999. Structure and Mechanism 

in Protein Science. New York: Freeman 

17. Gottesman S. 2004. The small RNA regulators 

of Escherichia coli: roles and 

mechanisms. Annu. Rev. Microbiol. 58: 

303–28 

18. Gromadski KB, Rodnina MV. 2004. Kinetic 

determinants of high-fidelity tRNA 

discrimination on the ribosome. Mol. Cell 

13:191–200 

19. Guerrier-Takada C, Gardinier K, Pace T, 

Altman S. 1983. The RNA moiety of ribonuclease 

P is the catalytic subunit of the 

enzyme. Cell 35:849–57 

20. Ha T. 2004. Structural dynamics and processing 

of nucleic acids revealed by sin- 


gle molecule spectroscopy. Biochemistry 

43:4055–63 

21. Ha T, Enderle T, Ogletree DF, Chemla 

DS, Selvin PR, Weiss S. 1996. Probing the 

interaction between two single molecules: 

fluorescence resonance energy transfer between 

a single donor and a single acceptor. 

Proc. Natl. Acad. Sci. USA 93:6264–68 

22. Ha T, Zhuang X, Kim H, Orr JW, 

Williamson JR, Chu S. 1999. Ligandinduced 

conformational changes observed 

in single RNA molecules. Proc. Natl. Acad. 

Sci. USA 96:9077–82 

23. Harlepp S, Marchal T, Robert J, Leger JF, 

Xayaphoummine A, et al. 2003. Probing 

complex RNA structures by mechanical 

force. Eur. Phys. J. E 12:605–15 

24. Heilman-Miller SL, Pan J, Thirumalai D, 

Woodson SA. 2001. Role of counterion 

condensation in folding of the Tetrahymena 

ribozyme II. Counterion-dependence 

of folding kinetics. J. Mol. Biol. 309:57– 

68 

25. Hess ST, Huang SH, Heikal AA, Webb 

WW. 2002. Biological and chemical applications 

of fluorescence correlation spectroscopy: 

a review. Biochemistry 41:697– 

705 

26. Jia YW, Sytnik A, Li LQ, Vladimirov S, 

Cooperman BS, Hochstrasser RM. 1997. 

Nonexponential kinetics of a single 

tRNA(Phe) molecule under physiological 

conditions. Proc. Natl. Acad. Sci. USA 

94:7932–36 

27. Kim HD, Nienhaus GU, Ha T, Orr 

JW, Williamson JR, Chu S. 2002. 

Mg 2+ -dependent conformational change of 

RNA studied by fluorescence correlation 

and FRET on immobilized single 

molecules. Proc. Natl. Acad. Sci. USA 99: 

4284–89 

28. Kruger K, Grabowski PJ, Zaug AJ, Sands 

J, Gottschling DE, Cech TR. 1982. Selfsplicing 

RNA: auto-excision and autocyclization 

of the ribosomal-RNA intervening 

sequence of Tetrahymena. Cell 

31:147–57 

29. Lindorff-Larsen K, Vendruscolo M, Paci





412 ZHUANG 

E, Dobson CM. 2004. Transition states for 

protein folding have native topologies despite 

high structural variability. Nat. Struct. 

Mol. Biol. 11:443–49 

30. Liphardt J, Dumont S, Smith SB, Tinoco I, 

Bustamante C. 2002. Equilibrium information 

from nonequilibrium measurements in 

an experimental test of Jarzynski’s equality. 


31. Liphardt J, Onoa B, Smith SB, Tinoco I, 

Bustamante C. 2001. Reversible unfolding 

of single RNA molecules by mechanical 

force. Science 292:733–37 

32. Lu HP, Xun LY, Xie XS. 1998. Singlemolecule 

enzymatic dynamics. Science 

282:1877–82 

33. Moerner WE, Orrit M. 1999. Illuminating 

single molecules in condensed matter. Science 

283:1670–76 

34. Mrksich M, Whitesides GM. 1997. Using 

self-assembled monolayers that present 

oligo(ethylene glycol) groups to control the 

interactions of proteins with surfaces. ACS 

Symp. Ser. 680:361–73 

35. Nahas MK, Wilson TJ, Hohng SC, Jarvie 

K, Lilley DMJ, Ha T. 2004. Observation of 

internal cleavage and ligation reactions of a 

ribozyme. Nat. Struct. Mol. Biol. 11:1107– 

13 

36. Nie SM, Zare RN. 1997. Optical detection 

of single molecules. Annu. Rev. Biophys. 

Biomol. Struct. 26:567–96 

37. Nissen P, Hansen J, Ban N, Moore PB, 

Steitz TA. 2000. The structural basis of ribosome 

activity in peptide bond synthesis. 


38. Noller HF, Hoffarth V, Zimniak L. 1992. 

Unusual resistance of peptidyl transferase 

to protein extraction procedure. Science 

265:1709–12 

39. Nugent-Glandorf L, Perkins TT. 2004. 

Measuring 0.1-nm motion in 1 ms in an 

optical microscope with differential backfocal-plane 

detection. Opt. Lett. 29:2611– 

13 

40. Onoa B, Dumont S, Liphardt J, Smith SB, 

Tinoco I, Bustamante C. 2003. Identifying 

kinetic barriers to mechanical unfold- 

ing of the T. thermophila ribozyme. Science 

299:1892–95 

41. Pan J, Deras ML, Woodson SA. 2000. Fast 

folding of a ribozyme by stabilizing core 

interactions: evidence for multiple folding 

pathways in RNA. J. Mol. Biol. 296:133– 

44 

42. Pan J, Thirumalai D, Woodson SA. 1997. 

Folding of RNA involves parallel pathways. 

J. Mol. Biol. 273:7–13 

43. Pan J, Woodson S. 1998. Folding intermediates 

of a self-splicing RNA: mis-pairing 

of the catalytic core. J. Mol. Biol. 280:597– 

609 

44. Pan J, Woodson SA. 1999. The effect of 

long-range loop-loop interactions on folding 

of the Tetrahymena self-splicing RNA. 

J. Mol. Biol. 294:955–65 

45. Pan T, Fang X, Sosnick TR. 1999. Pathway 

modulation, circular permutation and rapid 

folding under kinetic control. J. Mol. Biol. 

286:721–31 

46. Pan T, Sosnick TR. 1997. Intermediates 

and kinetics traps in the folding of a large 

ribozyme revealed by circular dichroism 

and UV absorbance spectroscopies and catalytic 

activity. Nat. Struct. Biol. 4:931– 

38 

47. Pljevaljcic G, Millar DP, Deniz AA. 2004. 

Freely diffusing single hairpin ribozymes 

provide insights into the role of secondary 

structure and partially folded states in RNA 

folding. Biophys. J. 87:457–67 

48. Rhoades E, Gussakovsky E, Haran G. 2003. 

Watching protein folding one molecule at a 

time. Proc. Natl. Acad. Sci. USA 100:3197– 

203 

49. Rodnina MV, Wintermeyer W. 2001. Fidelity 

of aminoacyl-tRNA selection on the 

ribosome: kinetic and structural mechanisms. 

Annu. Rev. Biochem. 70:415–35 

50. Rook MS, Treiber DK, Williamson JR. 

1998. Fast folding mutants of the Tetrahymena 

group I ribozyme reveal a rugged 

folding energy landscape. J. Mol. Biol. 

281:609–20 

51. Rueda D, Bokinsky G, Rhodes MM, 

Rust MJ, Zhuang X, Walter NG. 2004.





Single-molecule enzymology of RNA: essential 

functional groups impact catalysis 

from a distance. Proc. Natl. Acad. Sci. USA 

101:10066–71 

52. Russell R, Herschlag D. 1999. New pathways 

in folding of the Tetrahymena group 

IRNA enzyme. J. Mol. Biol. 291:1155–67 

53. Russell R, Millett IS, Tate MW, Kwok LW, 

Nakatani B, et al. 2002. Rapid compaction 

during RNA folding. Proc. Natl. Acad. Sci. 

USA 99:4266–71 

54. Russell R, Zhuang X, Babcock H, Millett 

IS, Doniach S, et al. 2002. Exploring 

the folding landscape of a structured RNA. 

Proc. Natl. Acad. Sci. USA 99:155–60 

55. Schutz GJ, Trabesinger W, Schmidt T. 

1998. Direct observation of ligand colocalization 

on individual receptor molecules. 

Biophys. J. 74:2223–26 

56. Sclavi B, Sullivan M, Chance MR, 

Brenowitz M, Woodson SA. 1998. RNA 

folding at millisecond intervals by synchrotron 

hydroxyl radical footprinting. Science 

279:1940–43 

57. Selvin PR. 1995. Fluorescence resonance 

energy transfer. Methods Enzymol. 

246:6264–68 

58. Shaevitz JW, Abbondanzieri EA, Landick 

R, Block SM. 2003. Backtracking by single 

RNA polymerase molecules observed at 

near-base-pair resolution. Nature 426:684– 

87 

59. Shav-Tal Y, Darzacq X, Shenoy SM, Fusco 

D, Janicki SM, et al. 2004. Dynamics of single 

mRNPs in nuclei of living cells. Science 

304:1797–800 

60. Silverman SK. 2003. Rube Goldberg 

goes (ribo)nuclear? Molecular switches 

and sensors made from RNA. RNA 9:377– 

83 

61. Strick TR, Croquette V, Bensimon D. 2000. 

Single-molecule analysis of DNA uncoiling 

by a type II topoisomerase. Nature 

404:901–4 

62. Strick TR, Kawaguchi T, Hirano T. 2004. 

Real-time detection of single-molecule 

DNA compaction by condensin I. Curr. 

Biol. 14:874–80 


63. Stryer L, Haugland RP. 1967. Energy transfer: 

a spectroscopic ruler. Proc. Natl. Acad. 

Sci. USA 58:719–26 

64. Sullenger BA, Gilboa E. 2002. Emerging 

clinical applications of RNA. Nature 

418:252–58 

65. Sytnik A, Vladimirov S, Jia YW, Li LQ, 

Cooperman BS, Hochstrasser RM. 1999. 

Peptidyl transferase center activity observed 

in single ribosomes. J. Mol. Biol. 

285:49–54 

66. Takamoto K, He Q, Morris S, Chance 

MR, Brenowitz M. 2002. Monovalent 

cations mediate formation of native tertiary 

structures of the Tetrahymena thermophila 

ribozyme. Nat. Struct. Biol. 9: 

928–33 

67. Tan E, Wilson TJ, Nahas MK, Clegg RM, 

Lilley DM, HA T. 2003. A four-way junction 

accelerates hairpin ribozyme folding 

via a discrete intermediate. Proc. Natl. 

Acad. Sci. USA 100:9308–13 

68. Tinoco I. 2004. Force as a useful variable in 

reaction: unfolding RNA. Annu. Rev. Biophys. 

Biomol. Struct. 33:363–85 

69. Treiber DK, Rook MS, Zarrinkar PP, 

Williamson JR. 1998. Kinetic intermediates 

trapped by native interactions in RNA 

folding. Science 279:1943–46 

70. Valadkhan S, Manley JL. 2002. Intrinsic 

metal binding by a spliceosomal RNA. Nat. 

Struct. Biol. 9:498–99 

71. Villa T, Pleiss JA, Guthrie C. 2002. Spliceosomal 

snRNAs: Mg2 + -dependent chemistry 

at the catalytic core? Cell 109:149–52 

72. Walter NG, Burke JM. 1997. Real-time 

monitoring of hairpin ribozyme kinetics 

through base-specific quenching of 

fluorescein-labeled substrates. RNA 3:392– 

404 

73. Walter NG, Burke JM. 1998. The hairpin 

ribozyme: structure, assembly and catalysis. 

Curr. Opin. Chem. Biol. 2:24–30 

74. Wassarman KM. 2004. RNA regulators of 

transcription. Nat. Struct. Mol. Biol 11: 

803–4 

75. Weiss S. 2000. Measuring conformational 

dynamics of biomolecules by single





414 ZHUANG 

molecule fluorescence spectroscopy. Nat. 

Struct. Biol. 7:724–29 

76. Widengren J, Rigler R. 1998. Fluorescence 

correlation spectroscopy as a tool to 

investigate chemical reactions in solutions 

and on cell surfaces. Cell. Mol. Biol. 

44:857–79 

77. Xie XS, Trautman JK. 1998. Optical studies 

of single molecules at room temperature. 

Annu. Rev. Phys. Chem. 49:441–80 

78. Xie Z, Srividya N, Sosnick TR, Pan T, 

Scherer NF. 2004. Single-molecule studies 

highlight conformational heterogeneity in 

the early folding steps of a large ribozyme. 

Proc. Natl. Acad. Sci. USA 101:534– 

39 

79. Zarrinkar PP, Wang J, Williamson JR. 

1996. Slow folding kinetic of RNase P 

RNA. RNA 2:564–73 

80. Zarrinkar PP, Williamson JR. 1994. Kinetic 

intermediates in RNA folding. Science 

265:918–24 

81. Zhuang X, Bartley L, Babcock H, Russell 

R, Ha T, et al. 2000. A single 

molecule study of RNA catalysis 

and folding. Science 288:2048–51 

82. Zhuang X, Kim H, Pereira M, Babcock 

H, Walter N, Chu S. 2002. Correlating 

of structural dynamics and function in 

single ribozyme molecules. Science 296: 

1473–77 

83. Zhuang X, Rief M. 2003. Single-molecule 

folding. Curr. Opin. Struct. Biol. 13: 

88–97



Zhuang.qxd 2/10/05 1:02 PM Page 1 

SINGLE-MOLECULE RNA SCIENCE C-1 

Figure 1 A single-molecule FRET study of the Tetrahymena ribozyme. (a) The 

Tetrahymena ribozyme. The Cy3 and Cy5 dyes serve as the fluorescent donor and 

acceptor. The ribozyme was immobilized to surface via a biotin-streptavidin interaction. 

(b) Folding time trajectories showing the unfolding and folding of a single 

ribozyme molecule. Red trace: the acceptor signal; green trace: the donor signal; 

blue trace: the FRET value, defined as the acceptor signal divided by the sum of the 

donor and acceptor signals. The unfolding and refolding were triggered by removing 

and adding Mg 2+ at 3 s and 30 s, respectively. Figure adopted from Reference 81.




C-2 ZHUANG 

Figure 2 A single-molecule FRET study of the hairpin ribozyme. (a) The two-way junction 

hairpin ribozyme. The Cy3 and Cy5 dyes serve as the fluorescent donor and acceptor. 

(b) The FRET time traces of individual ribozyme molecules show heterogeneous undocking 

kinetics (upper three panels). The FRET levels at 0.8 and 0.2 correspond to the docked 

and undocked states, respectively. The distributions of dwell times of undocked and docked 

states, obtained from hundreds of time traces, indicate that the two-way junction ribozyme 

docks with a single rate constant but undocks with at least four different rate constants 

given in (c). Conversion between different undocking behaviors requires at least several 

hours (lower panel). (c) The reaction pathway of the hairpin ribozyme. Rz, ribozyme; S, 

substrate; 3�P, 3� cleavage product; 5�P, 5� cleavage product. The rate constants were 

obtained from single-molecule FRET experiments in conjunction with ensemble cleavage 

and ligation assays. Figure adopted from Reference 82.




SINGLE-MOLECULE RNA SCIENCE C-3 

Figure 3 A model of tRNA accommodation into the A site of the ribozyme inferred from a recent single-molecule FRET experiment and 

previous kinetic characterizations. The ribozyme is shown in yellow. Exit (E), peptidyl (P), and aminoacyl (A) sites are depicted as yellow 

rectangles. tRNAs are shown in blue. The elongation factor Tu (EF-Tu) is shown in three colors (light blue, red, and purple), each corresponding 

to a different state. Step 1: Initial binding of the EF-Tu(GTP)tRNA ternary complex with the ribosome. Step 2: Contact is made 

between the anticodon of tRNA and the messenger RNA codon. Step 3: Productive codon recognition triggers the tRNA to move closer to 

the P site. Step 4: GTP hydrolysis. Step 5: EF-Tu changes from the GTP- to the GDP-bound form. Step 6: Release of tRNA from EF-Tu 

to accommodate at the peptidyl transferase center. Step 7: Peptide-bond formation. The value of FRET between the two tRNAs is indicated. 

Figure adopted from Reference 5.




C-4 ZHUANG 

Figure 4 A single-molecule force study of the Tetrahymena ribozyme. (a) Secondary 

structure of the Tetrahymena ribozyme. Gray lines label sequences targeted by complementary 

DNA oligonucleotides, and “M” labels are site-directed mutations, both of which 

were used for identifying the molecular interactions disrupted at each unfolding step (kinetic 

barriers for mechanical unfolding). The letters a through h indicate the proposed 

positions of the kinetic barriers on the ribozyme. (b) Representative unfolding (black) 

and refolding (pink) force-extension curves of the ribozyme displaying several unfolding 

steps. The letters a through h correspond to the kinetic barriers. Figure adopted from 

Reference 40.

Single-Molecule RNA Science - Xiaowei Zhuang - Harvard University

You also want an ePaper? Increase the reach of your titles

Delete template?

Save as template?