Nature Reviews Molecular Cell Biology 4, SS1–SS5 (2003)

Review: Single-molecule visualization in cell biology

Recent progress in single-molecule detection techniques has allowed us to visualize the dynamic behaviour and reaction kinetics of individual biological molecules inside living cells. Single-molecule visualization provides a direct way to quantify, with a high spatial and temporal resolution, biological events inside cells at the single-molecule level. In this article, we discuss how single-molecule visualization can be used in cell biology.

Yasushi Sako^1,2 and Toshio Yanagida^1,3
1Nanobiology Laboratories, Graduate School of Frontier Biosciences, Osaka University, 2-2 Yamadaoka, Suita, Osaka 565-0871, Japan.
²Time's Arrow and Biosignaling, PRESTO, Japan Science and Technology Corporation, Japan.
³Soft Nanomachine Program, Japan Science and Technology Corporation, 2-2 Yamadaoka, Suita, Osaka 565-0871, Japan.
correspondence to: sako@phys1.med.osaka-u.ac.jp

Published online: 1 September 2003
doi:10.1038/nrm1193

Single-molecule analysis is a powerful method to study purified biomolecules in vitro, because the data obtained are not obscured by the averaging that is inherent in conventional biochemical experiments^1,2,3. Recently, this advantage has been extended to studies using living cells^4,5, in which it has been possible to quantify the dynamic and kinetic parameters of single-molecule reactions in vivo. It is difficult to determine kinetic parameters using multiple-molecule techniques because the reactions of individual molecules occur stochastically inside a cell. In addition, it is hard to spot local and temporal heterogeneities in the dynamic movement of molecules using multiple-molecule techniques. Single-molecule techniques can therefore be used to avoid such difficulties. In this review, we illustrate how single fluorescent molecules can be visualized in living cells. We then discuss the unique and important findings in cell biology that could only have been obtained using single-molecule visualization, as well as the recent progress in in vitro measurements that combine single-molecule visualization with other single-molecule techniques.

Why single-molecule visualization?

Single-molecule visualization of fluorophores in aqueous conditions — using TOTAL INTERNAL REFLECTION fluorescence microscopy (TIR-FM)⁶ or epi-fluorescence microscopy^6,7 — was first performed in 1995. Since then, this technique has been applied to observe directly the motions of linear and rotational molecular motors, enzymatic reactions, the structural dynamics of proteins and DNA–protein interactions in vitro^1,2,3. The visualization of single lipid molecules in a lipid bilayer and the simultaneous measurement of ion conductance and the movement of single ion channels in a model membrane have also been reported^2,4. Furthermore, in 2000, the biological reactions of ligand and receptor proteins⁸ and the movements of lipid molecules⁹ were first visualized as single molecules on the surface of living cells (see Movie 1 online). Since then, ion channels¹⁰, small G proteins and their effectors¹¹, cell-adhesion proteins¹², viral proteins¹³ and components of the cytoskeleton¹⁴ have been visualized as single molecules in living cells. The list of molecules that have been visualized is still growing rapidly, and the results of some of these studies are discussed in more detail below.

The advantages of single-molecule analyses. Proteins in living cells work as part of molecular networks that have specific functions, such as gene expression, energy transduction or membrane transport. Advances in molecular biology are rapidly uncovering the components of these molecular networks and are determining how they are constructed. One of the next objectives of cell biology is to quantify the flow of materials, information and energy through these molecular networks. To achieve this, both the dynamic and kinetic parameters of the single processes within the networks — such as the movement and translocation of proteins, the protein–protein interactions and the enzymatic reactions — must be determined in living cells. Single-molecule analysis in vivo will prove to be a powerful technique for this purpose. This technique has the ultimate level of sensitivity and can be used to detect and observe single reactions in living cells. In addition, it provides several other important advantages.

First, reactions do not need to be synchronized for single-molecule measurements. Statistically, each protein molecule reacts at a different time and position in a cell, so, to obtain the dynamic and kinetic parameters of a reaction using multiple-molecule measurements, specific techniques are required that synchronize the start-point of the reaction of every molecule. However, it is not possible to synchronize the intermediate steps of a reaction network, and single-molecule measurements have the advantage of bypassing these problems.

Second, single-molecule measurements provide information about the fluctuations and distributions of dynamic and kinetic parameters. Such information cannot be obtained from ensemble averages. Considering the local heterogeneity in the structure and environment of living cells, the fluctuation and distribution of the reactions of each molecule are probably important in understanding the mechanisms of cellular events.

Third, the above two points implicitly mean that single-molecule analysis allows the relationship between the inputs and outputs of single events of protein reactions to be quantified. In single-molecule experiments, each input and output event — for example, the binding and dissociation of a ligand or substrate — can be monitored individually. To understand reaction cascades in cells, the relationship between all the inputs and outputs of every step must be understood.

Single-molecule analysis requires statistical data so that the observed behaviour of minor, unusual molecules is not overestimated. However, monitoring many single molecules for statistical analysis is a laborious task. This is because automatic image processing is difficult for single-molecule experiments in living cells due to a limited signal-to-noise ratio and a non-homogeneous background. In addition, as signals cannot be obtained from invisible molecules, the appropriate controls, which depend on the purpose of the experiment, should be used. Despite these drawbacks, single-molecule studies provide unique insights in cell biology and so are extremely useful. As single-molecule studies deal with small numbers of molecules, sampling noise is an inevitable problem of these analyses compared with conventional biochemical analyses. However, this high level of sampling noise might be an essential factor in cell behaviour because the number of protein molecules per cell is usually relatively small (for example, most cell-signalling proteins are present at a concentration of 10²–10⁵ molecules per cell).

Techniques for single-molecule visualization. TIR-FM is a widely used technique for single-molecule detection both in vitro and in vivo^2,5. TIR-FM, which was originally developed to observe the interface between two media with different diffractive indices¹⁵, uses an electromagnetic field called the 'evanescent field' to excite fluorophores (Fig. 1). As the evanescent field diminishes exponentially with distance from the interface, the excitation depth in TIR-FM is limited to a very narrow range — typically one hundred to several hundreds of nanometres. However, using such a narrow excitation depth is the most effective way to overcome the background noise problem, which is often the greatest problem of single-molecule imaging.

Fig. 1
		Figure 1 | Total internal reflection fluorescence microscopy.

Objective-type TIR-FM¹⁶, in which the excitation laser beam illuminates the specimen through an objective lens, is particularly useful for imaging living cells (Fig. 1). The top surface of the specimen is free in this type of TIR-FM, so it can be combined with high-resolution differential interference contrast microscopy, which requires an oil immersion condenser, and allows the cells to be easily accessed for changes of the surrounding medium, microinjection or micromanipulation. Using objective-type TIR-FM, precise morphological information about the cells can be obtained, the effect of drugs can be examined and electrophysiological experiments can be performed at the same time as single-molecule visualization.

Although TIR-FM provides superior contrast compared with other far-field microscopy techniques, its application is limited to the proximity of the cell surface — that is, to studying parameters in two dimensions. To observe single molecules deep inside cells in three dimensions, conventional epi-fluorescence microscopy using a laser for excitation⁹ and real-time confocal microscopy¹⁷ are applicable. The latter is thought to produce better results than TIR-FM for single-molecule imaging in dense solutions. Only sparsely labelled samples (<10 particles/µm²) can be visualized as single molecules using TIR-FM, epi-fluorescence microscopy or confocal fluorescence microscopy owing to the low spatial resolution. However, the higher spatial resolution of SCANNING NEAR-FIELD OPTICAL MICROSCOPY might be able to overcome this limitation in the future¹⁸. Single molecules that are rapidly moving in solution cannot be visualized as fluorescent spots owing to the rapid three-dimensional Brownian diffusion that occurs. However, FLUORESCENCE CORRELATION SPECTROSCOPY¹⁹, which allows single molecules to be analysed in solution, can be used to complement single-molecule visualization.

Applications in living cells

Single-molecule analyses have been effectively used both to quantify intracellular reactions and for the spatial analysis of such reactions with high resolution: for example, the kinetic analysis of receptor–ligand interactions²⁰; measuring protein dynamics in the cytoplasm²¹, nucleus²² or plasma membrane^9,10,13,23; observing the assembly and disassembly of protein oligomers^8,12 or large protein complexes¹⁴; and detecting the chemical reactions of proteins⁸. Examples of such studies will be discussed in more detail below.

Kinetic analysis of receptor–ligand interactions. The dissociation kinetics of a receptor–ligand interaction is an important property of cell-signalling reactions. Dictyostelium discoideum amoebae perform chemotaxis towards cyclic AMP, and this chemotaxis is induced through a G-protein-coupled receptor²⁴. To determine how cells sense gradients of cAMP, the binding of fluorophore-labelled cAMP (Cy3–cAMP) to its receptor was observed on the surface of cells¹¹. The dissociation rate constant for cAMP from its receptor was obtained by performing a statistical analysis of the time between the binding and dissociation of single molecules of Cy3–cAMP on the cells undergoing chemotactic movements²⁰ (Fig. 2a).

Fig. 2
		Figure 2 | Single-molecule analysis of cellular events.

Cy3–cAMP dissociated faster from the anterior half of the cells compared to the posterior half, although the density of cAMP binding was almost uniform over the entire cell surface. This indicates that the association–dissociation cycle of cAMP is faster at the anterior end of the cell. Therefore, the reaction state of the cAMP receptors depends on the location of the chemotactic cells in relation to the concentration gradient of the cAMP signalling molecule (that is, the cAMP concentration gradient is converted to the difference in the reaction state of the cAMP receptor). This difference seems to depend on the coupling between the cAMP receptor and the trimeric G protein on the cytoplasmic side of the plasma membrane, and it is highly probable that cAMP signalling is more active in the anterior region of chemotactic amoeba than in the posterior region.

The dynamics of membrane components. It is possible to use single-molecule imaging to probe the dynamic and microscopic structures of the plasma membrane by measuring the movement of the membrane components (Fig. 2b). For example, the lateral diffusion of fluorescent lipids that were incorporated into a muscle-cell membrane has been observed for individual molecules⁹. In this study, a lipid probe with saturated acyl chains was found to be confined to small regions of the plasma membrane (spanning 0.7 µm), which indicates the presence of small lipid microdomains that are probably related to membrane rafts. In addition, the study also detected the shape and motions of the microdomains using single-molecule microscopy. As membrane rafts contain various cell-signalling molecules, the movement of the raft components are likely to be important for signal transduction from the plasma membrane.

Using a similar technique, single-molecule imaging of ion channels in the plasma membrane has been performed^10,23, and the data indicate that different types of ion channel behave in distinct ways. For example, a T-lymphocyte K⁺ channel that was studied by single-molecule imaging presented as monomers and was essentially immobile¹⁰, whereas a human cardiac Ca²⁺ channel tended to form larger aggregates and was mobile²³. Virus entry into a living cell has also been observed by visualizing the dynamics of single viral proteins¹³, and this single-molecule detection has a much higher sensitivity compared to conventional methods that require the accumulation of many viral particles in a cell. These examples highlight the potential of single-molecule techniques as new and sensitive assays in pharmacology and pathology.

Dynamics of cytoskeletal filaments. Single fluorescent tracers that are incorporated into large supramolecules can be used to monitor the dynamics of entire complexes. Watanabe and Mitchison¹⁴ have used single molecules of actin that are tagged with green fluorescent protein (GFP–actin) as a tracer for analysing the spatial regulation of actin-filament dynamics in live fibroblast cells (Fig. 2c). The expression of GFP–actin was ingeniously controlled by truncating the enhancer region of the expression plasmid to achieve a diluted labelling of endogenous actin (1 GFP–actin molecule to 10,000–50,000 endogenous actin molecules). Incorporation into an actin filament, retrograde movement towards the cell centre and detachment from the filament was then observed for individual molecules of GFP–actin. There has been a long-standing controversy about the position of actin polymerization along an actin filament, and the appearance and duration of single fluorescent spots of GFP–actin that were incorporated into actin filaments allowed the kinetic parameters of polymerization and depolymerization to be determined. Modelling of the results indicates that basal polymerization and depolymerization are constant throughout lamellipodia — flattened, sheet-like structures that project from the surface of a cell and are composed of a meshwork of crosslinked F-actin — and that most of the actin filaments in the lamellipodia are generated by this basal polymerization. Additional polymerization within 1 µm of the tip of the cell is thought to balance the movement of actin filaments towards the centre of the cell.

Monitoring chemical reactions. Monitoring chemical reactions will be an important future goal of single-molecule analysis in living cells, and a method has been developed to detect the tyrosine phosphorylation of a membrane receptor in terms of single molecules⁸ (Fig. 2d). Single-molecule analysis^8,11 has confirmed that epidermal growth factor (EGF) binding to the EGF receptor (EGFR) induces the dimerization of EGFR, which, in turn, is followed by autophosphorylation of the cytoplasmic tyrosine residues of the EGFR²⁵.

In these experiments^8,11, a monoclonal antibody that recognizes the cytoplasmic domain of phosphorylated EGFR was labelled with the fluorophore Cy3 and was used to detect the activation of the EGFR. EGF was conjugated to another fluorophore (Cy5). To introduce the Cy3–antibody into the cytoplasm, semi-intact cells that were perforated by streptolysin O were used²⁶. After pulse stimulation of the semi-intact cells by the addition of Cy5–EGF, Cy3–antibody and ATP were added and the binding of Cy5–EGF and Cy3–antibody to the plasma membrane were simultaneously visualized as single molecules. Fluorescent spots of Cy3–antibody tended to co-localize with the clusters of Cy5–EGF, which indicates autophosphorylation of the EGFR clusters that contain bound EGF. So, using this technique, a widely accepted hypothesis for the mechanism of EGFR activation — that is, that the formation of EGF–EGFR dimers is necessary for the autophosphorylation of the EGFR — was proved directly.

Single-molecule analysis in vitro

From the work discussed so far, it is clear that single-molecule visualization in vivo can be used to detect the positions and movements of individual single molecules, or to detect co-localization between two or more single molecules. SINGLE-PAIR FLUORESCENCE RESONANCE ENERGY TRANSFER¹ can be used to detect intermolecular structural change in vitro⁸. In addition, single-molecule FLUORESCENCE POLARIZATION and spectroscopy can be used to detect the movements and intramolecular structural changes of single molecules and the microenvironment around single molecules. However, the functions of molecules usually cannot be assessed directly by these techniques. To allow single-molecule analysis to evolve further, techniques that visualize single molecules must be combined with techniques that can detect the functions of these molecules. In fact, such combinations have already been successfully achieved in some recent experiments in vitro.

Combination with single-molecule manipulation: analysis of molecular motors. Biological molecules can be manipulated directly using a glass needle or the tip of a cantilever of an ATOMIC-FORCE MICROSCOPE or, less directly, using beads that are trapped by OPTICAL TWEEZERS²⁷. Using these techniques, the mechanical properties of single biological molecules can be determined to nanometre and piconewton levels of accuracy for displacement and force, respectively.

For example, a molecular motor, kinesin, has been found to move processively along a microtubule with regular 8-nm steps, which indicates that kinesin 'walks' along the – tubulin dimer repeat²⁸. In addition, the step size of muscle myosin (myosin-II) has been determined to be 5–15 nm, although this step size has not always been consistent among all researchers (for reviews, see Refs 29–31). To understand how molecular motors work, it is crucial to know how the mechanical events are coupled to the ATP hydrolysis event. Many techniques have provided insights and, by combining the nano-manipulation technique with the single-molecule imaging technique, we have been able to observe the coupling between individual mechanical and chemical (that is, ATP hydrolysis) events of single muscle myosin molecules directly³². The next key question is how the power stroke of the molecular motor is developed. There is a large body of evidence that indicates conformational changes in the neck domain of myosin-II heads. Based on these findings, a lever-arm model is proposed for the movement of muscle myosin and, in this model, the neck domain acts as a lever arm, the tilting of which causes displacement of myosin³³. This model predicts that the step size is proportional to the length of the neck domain.

Single-molecule imaging and manipulation techniques have also been used to study unconventional myosins — members of the myosin superfamily that are more amenable to movement studies than conventional myosins. However, these myosins are structurally different to conventional myosins and are not found in muscle. Their physiological roles are often unknown or unclear, although they are known to travel long distances continuously along actin filaments to transport cellular cargo (Fig. 3a,b). A class-V myosin (myosin-V) with long neck domains moves processively along actin filaments with large steps^34,35 that alter the angle of a fluorophore attached to the neck domain before and after stepping³⁶. These results are consistent with a lever-arm model for myosin-V movement, although they do not prove directly that it is the tilting of the neck domain that is causing the steps. However, data have been obtained that question the lever-arm model for the movement of unconventional myosins^37,38,39 and, on the basis of these data, we have suggested that another mechanism, such as the biased linear diffusion of myosin heads along an actin filament, might be operating for the processive movement of these myosins⁴⁰ (see Movie 2 online).

Fig. 3
		Figure 3 | Combinations of single-molecule techniques for in vitro protein analysis.

Combination with single-molecule electrophysiology: analysis of channel proteins. The development of technology to record the current through a single ion channel allowed the functions of single proteins to be directly assessed both in vitro and in vivo⁴¹. This technology has shed light on the kinetic and pharmacological properties of many kinds of ion channel, although detailed mechanisms of ion-channel function are still being determined. The ability to monitor the conformation and chemical state of ion channels, combined with measurements of single-channel ion currents, would aid research in this area.

As mentioned earlier, single ion channels have been visualized in living cells^10,23 and, recently, Sonnleitner et al. visualized single molecules of a voltage-gated K⁺ channel conjugated to tetramethylrhodamine in living cells⁴². In this study, the fluorescent intensity changed at the same time as the membrane potential, which indicates that rearrangements of the protein structure were being detected. By combining artificial-lipid-bilayer and single-molecule-imaging techniques, an experimental system has been developed that has allowed single ion-channel currents and single-molecule images of ion-channel molecules to be obtained simultaneously⁴³ (Fig. 3c,d). Therefore, it should not be long before simultaneous observation of the conformation and chemical states of single ion channels together with single-molecule ion currents is possible, both in artificial membranes and on the surface of living cells.

Conclusion and perspectives

Developments in single-molecule techniques in living cells have allowed us to visualize the location and movements of molecules and to determine the number of molecules that are involved in cellular reactions. Reaction kinetics of intracellular molecules have been measured and the activation of molecules has been detected. Single-molecule visualization in living cells has proven useful for quantifying cellular reactions and will be indispensable for further understanding the molecular mechanisms of cellular responses. Single-molecule techniques can be used at two levels — monitoring cellular responses in terms of the reactions of single-molecules and elucidating the mechanisms of how proteins work in terms of dynamic, kinetic and conformational changes of single molecules. Single-molecule manipulation using fine glass needles, optical tweezers or atomic-force microscopy has become widely used for protein studies in vivo. The manipulation of membrane proteins at the single-molecule level has already been reported in living cells⁴⁴, and 'single-channel recording'⁴¹ is another technique that is used to assess the functions of single molecules in living cells. In the future, combining single-molecule-visualization, single-molecule-manipulation and single-molecule-electrophysiology techniques will be important to allow us to further understand the nanobiology of living cells.

Acknowledgements

We thank the former members of the Single Molecule Processes Project, Japan Science and Technology Corporation (JST), and the members of the Nanobiology Laboratories of Osaka University, Japan, for their cooperation.

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