Dynamics have been traditionally measured using bulk methods, with signals sensitive to dynamic changes for a large collection of molecules. These signals, such as emission from a fluorescent dye, must be sensitive to conformational or compositional changes of the system as it evolves in time. The problem arises in synchronizing a large collection of molecules to detect a change in a bulk signal. Imagine a fluorescence signal that changes upon tRNA binding to the A-site (this could be from a dye on a tRNA that yields different emission probabilities whether on a free or ribosome-bound tRNA). In order to detect a change in this signal, the system must be synchronized such that all tRNAs are unbound at the start of the measurement, usually through rapid mixing. This ensures that the observed time-dependent fluorescence change reports only on the approach to equilibrium from the unbound state, and from this signal kinetic information can be extracted. However, if we want to look at a subsequent tRNA binding event, we would have to pause the evolution of the system, remix the reagents, and repeat the measurement. While rapid double-mixing experiments are possible with existing instrumentation, these have limited applicability to the ribosome and are not extensible to the observation of many iterations of peptide bond formation. In short, dynamics cannot be measured in real time during multiple rounds of elongation. This need for synchronization is a fatal limitation of bulk kinetic investigations to probe dynamics.
Single-molecule experiments allow direct measurement of dynamics without the need for synchronization. Organic fluorophores emit sufficient numbers of photons to be readily detected with modern cameras at millisecond time resolution. To distinguish weak single-fluorophore fluorescence from background illumination and noise, various techniques such as prism-based total internal reflection fluorescence microscopy (TIRF), objective-based TIRF and zero mode waveguides (ZMWs) are utilized to constrain the volume that is illuminated, which drastically reduces background fluorescence. The photophysical stability of the fluorophores depends on the illumination intensity and reaction conditions and ranges from seconds to tens of minutes. Thus, signals from individual molecules in principle can be observed directly over a period of time from milliseconds to hours. However, the rapid diffusion of the fluorophores in solution limits observation of the free molecules, since on the timescale of data acquisition a labeled molecule leaves the illumination volume and is delocalized, not producing enough photons to be observed along its diffusion track (r.m.s. diffusion distances for tRNA and the 70S ribosome are ~ 1.3 and 0.5 µm in 10 ms respectively).
These problems are turned into advantages by spatially constraining the system through surface immobilization – in translation experiments this is most often accomplished using biotin-streptavidin interactions to bind mRNAs or ribosomes to an optically transparent surface. Let’s take a simple example of the bimolecular binding event of a dye-labeled tRNA to an immobilized ribosome. In the single-molecule fluorescence experiment, the freely diffusing unbound tRNA is invisible and binding of the tRNA to an immobilized ribosome leads to a burst of observed fluorescence, as the fluorophore, emitting a large number of photons, is now localized within a small observation volume, as opposed to freely diffusing in solution. Binding of multiple tRNAs is manifested at the single-molecule level as a series of fluorescence bursts and inter-burst delays. The statistics of the burst lifetimes and delay times are exponentially distributed for a simple two-step process, yielding time constants that are reciprocal to the rate constants for dissociation and association, respectively.
The power of this approach is revealed when we look at binding of a second tRNA labeled with a differently colored dye. The single-molecule analysis of this multi-color experiment would reveal the relative arrival time for the two different tRNAs, by the time interval between the fluorescence bursts. Further analysis would also reveal how long the two bound tRNAs overlapped on the same ribosome. These results can be obtained directly from the data without the need to synchronize the ribosomes experimentally. The subsequent single-molecule traces can be analyzed in a variety of ways to extract dynamic data. “Post-synchronization” in silico extracts the relative timing of two or more events. For example, statistics of arrival order, or the average overlap time of occupancy by two tRNAs or other ligands can be calculated. Thus, correlations between dynamic events are observed directly.
Conformational dynamics are readily investigated using single-molecule fluorescence. The main tool for this application is Förster resonance energy transfer (FRET), which involves energy transfer between a donor and acceptor dye through coupling of transition dipoles. The efficiency of energy transfer depends on 1/R6 where R is the inter-dye distance, as well as on the spectral overlap of the two dyes and on dye orientation terms. For standard dyes used in these experiments (Cy3 and Cy5), the efficiency of FRET varies from 1 (all emissive energy from Cy3 is transferred to Cy5) at distances below about 20 Å to 0 at distances above 80 Å. Thus FRET is highly sensitive to distance changes in the region of 30–60 Å, well suited for investigation of the ribosome (250Å diameter).
FRET represents a high time-resolution probe of biomolecular conformation. In single-molecule FRET (smFRET), donor and acceptor dye emission are measured simultaneously for individual molecules, and these intensities converted to FRET through the equation: Eobs = Iacceptor/(Idonor + Iacceptor), where Idonor and Iacceptor are fluorescence intensities of the donor and acceptor dyes correspondingly. Changes in inter-dye distance are revealed by anticorrelated changes in donor and acceptor intensities. Molecular conformation can thus be monitored by smFRET with millisecond time resolution, limited by fluorophore brightness and camera sensitivity. smFRET has been a powerful tool to explore ribosomal and ligand dynamics during translation, as outlined below.
In addition to dynamics, single-molecule methods can directly measure or manipulate the forces generated by molecular motors. Single-molecule force methods employ optical tweezers to apply a mechanical force to biomolecular complexes and study their behavior under stress. Individual molecules are too small to permit precise nanomanipulations, such as moving, stretching, and so on. To that end, individual molecules are attached to two larger microscopic objects that could be more easily handled. Molecules are usually linked to a polystyrene bead that can be manipulated with an optical trap - a laser beam focused by high numerical aperture microscope objective. A polystyrene bead placed in the focus of the microscope objective is trapped via the radiative pressure gradient from a focused laser beam. Dislocation of the bead from the trap focus generates a restoring force proportional to the bead displacement. To allow application of tensile force, another end of the molecule is bound to the microscope slide surface or to the another bead that held by a second optical trap, thus stretching molecule between two points. Changing the relative position of the trap centers or moving the slide surface, allows control over the position of the bead relative to the center of the trap, allowing control over the applied force. Optical traps can apply constantly increasing force to the point of complex rupture to test mechanical stability of the complexes, thus directly reporting on the tensile strength of the intermolecular interactions, such as those between mRNA and the ribosome. Alternatively, the trap can be employed as a molecular ruler to track relative movement of the two components in the complex, for example the travelling of the ribosome along mRNA. Optical tweezers permit application of an intermediate assisting or hindering force to the molecular motor thus allowing elucidation of the mechanism of the motor mobility. Modern optical traps allow distance measurement at the Ångstrom level of resolution and application of forces in the range of tens of piconewtons, with sub-picowewton precision. The ribosome reads mRNA in 1.3 nm long triplets and requires up to 20 pN to dislocate from mRNA, thus it is suitable for study with single-molecule force methods.
Single-molecule experiments must be tackled with care and diligence. The approaches, by their very sensitivity, are fraught with potential artifacts. Dye or bead labeling of molecules can perturb conformation and block interactions, so the function of all labeled molecules must be validated by independent assays. Likewise, surface immobilization can perturb behavior of biological systems, and non-specific surface interactions of biomolecules can interfere with single-molecule measurements. Biologically-relevant signals are convoluted with photophysical and photochemical artifacts due to high-intensity illumination — blinking, photobleaching and photodamage limit single-molecule measurements and must be addressed by use of specific dyes, removal of molecular oxygen and addition of chemical agents to improve dye behavior. Subsequent single-molecule data analysis is time-consuming and will require improvements in unbiased techniques to extract true dynamic behavior. The challenging and time-consuming nature of the single-molecule approach limits throughput. As result, single-molecule experiments should be used in conjunction with conventional kinetic and molecular biology methods to address biological questions. Despite these limitations, the past decade has seen single-molecule data contributing deeply to our understanding of translation.