Nature Cell Biology 5, S7–S14 (2003)

Review: Photobleaching and photoactivation: following protein dynamics in living cells

Cell biology is being transformed by the use of fluorescent proteins as fusion tags to track protein behaviour in living cells. Here, we discuss the techniques of photobleaching and photoactivation, which can reveal the location and movement of proteins. Widespread applications of these fluorescent-based methods are revealing new aspects of protein dynamics and the biological processes that they regulate.

Jennifer Lippincott-Schwartz, Nihal Altan-Bonnet and George H. Patterson
Cell Biology and Metabolism Branch, National Institute of Child Health and Human Development, Building 18T/Room 101, National Institutes of Health, Bethesda, Maryland 20892, USA.
correspondence to: jlippin@helix.nih.gov

Published online: 1 September 2003
doi:10.1038/ncb1032

The discovery and development of fluorescent proteins from marine organisms are revolutionizing the study of cell behaviour by providing convenient markers for gene expression and protein targeting in intact cells and organisms (see also the review on page S1 of this supplement)^1,2. The most widely used of these fluorescent proteins — green fluorescent protein (GFP) from the jellyfish Aequorea victoria ³ — can be attached to virtually any protein of interest and still fold into a fluorescent molecule. The resulting GFP chimaera can be used to localize previously uncharacterized proteins⁴ or to visualize and track known proteins to further understand cellular events⁵.

The use of GFP as a minimally invasive tool for studying protein dynamics and function has been stimulated by the engineering of mutant GFPs with improved brightness, photostability and expression properties^2,6,7. Cells that express proteins tagged with these GFPs can be imaged with low light intensities over many hours and so can provide useful information about changes in the steady-state distribution of a protein over time. Time-lapse imaging alone, however, cannot reveal a protein's kinetic properties (for example, whether it is freely diffusing, bound to an immobile scaffold, or undergoing binding with and dissociation from other components). Yet, it is these kinetic properties that are arguably of most interest, as they underlie protein function within cells.

In this review, we discuss two techniques — photobleaching and photoactivation — that, when combined with time-lapse imaging, can uncover the kinetic properties of a protein by making its movement observable^{2,7,8,9,10,11}. Photobleaching — the photo-induced alteration of a fluorophore that extinguishes its fluorescence — accomplishes this through fluorescence depletion within a selected region. Photoactivation, on the other hand, works by converting molecules to a fluorescent state by using a brief pulse of high-intensity irradiation. After fluorescently highlighting specific populations of molecules by either method, the fluorescent molecules can be followed as they re-equilibrate in the cell. The extent and rate at which this occurs can be quantified and used with computer-modelling approaches to describe the kinetic parameters of a protein.

Photobleaching techniques

Fluorescence recovery after photobleaching. Developed over two decades ago to study the diffusive properties of molecules in living cells^{12,13,14,15,16,17}, fluorescence recovery after photobleaching (FRAP) has experienced a resurgence due to the introduction of GFP and the development of commercially available confocal-microscope-based photobleaching methods^8,11,18. In this technique, a region of interest is selectively photobleached with a high-intensity laser and the recovery that occurs as molecules move into the bleached region is monitored over time with low-intensity laser light (Fig. 1a). Depending on the protein studied, fluorescence recovery can result from protein diffusion, binding/dissociation or transport processes.

Fig. 1
		Figure 1 | Fluorescence recovery after photobleaching.

Analysis of fluorescence recovery can be used to determine the kinetic parameters of a protein, including its diffusion constant, mobile fraction, transport rate or binding/ dissociation rate from other proteins. In experiments in which the protein of interest moves freely, the fluorescence will recover to the initial prebleach value and the shape of the recovery curve can be described mathematically with a single component recovery (Fig. 1b, single)^19,20,21,22. Determining the effective diffusion coefficient (D_eff ) and mobile fraction (M_f ) of a protein from such data is relatively straightforward, given the previous analysis of FRAP kinetics¹² (for several recent reviews, see Refs 8,11,18,23). If the shape of the curve is complex (that is, it requires a multi-component diffusion equation^20,24,25), then multiple populations of the molecule with differing diffusion rates are present (Fig. 1b, complex). This can occur when a molecule undergoes binding and release from intracellular components or exists as a monomer and multimeric forms¹⁰. Alternatively, the protein might not be diffusing but might be undergoing movement driven by molecular motors or membrane tension flow. A simple test for determining whether a fluorescent protein moves by diffusive movement or facilitated transport is to vary the size of the bleached area or beam radius, . The recovery will change with an ² dependence for diffusive movement only²⁶. Accurate analysis of FRAP data requires that the bleach event is much shorter than the recovery time and preferably as short as possible. Moreover, the recovery event must be monitored until a recovery plateau is achieved, which is much greater than the half-time for recovery. See Table 1 for other FRAP considerations.

Performing FRAP. Until recently, carrying out FRAP required custom-built systems to perform the measurements. Development of FRAP methods for use on the laser-scanning confocal microscope has made this technique widely available. Images on the confocal microscope are obtained by scanning a focused laser beam across the specimen and recording the emitted fluorescence through a pinhole that is situated in front of the light detector. One way to photobleach using this system is to define a region-of-interest at the highest possible ZOOM, set the laser power to maximum, and set the laser ATTENUATION to zero. The high zoom increases the dwell time of the laser on the bleached region per line scan (laser intensity increases proportionally to the square of the zoom factor), which therefore greatly increases the radiation per area. But a more advanced method is to use an acousto-optical tunable filter (AOTF; available on more recent commercially available confocal microscopes), which allows rapid (microsecond to millisecond) attenuation of the laser as it scans a field. By allowing rapid switching between the bleaching and normal beam, the AOTF allows accurate measurements of diffusion rates in defined areas.

Use of an AOTF also enables users to photobleach virtually any pattern or shape. This allows FRAP studies to be done on organelles of complex shapes, allowing the lateral mobility of organelle-specific membrane and lumenal proteins to be investigated. Selective photobleaching on a confocal microscope also provides a method for analysing aspects of protein dynamics other than diffusion (including assembly/disassembly of protein complexes in cells, the exchange of cytosolic proteins on and off organelles, and the lifetime and fate of membrane-bound transport intermediates^27,28,29,30 (Fig. 2a). This type of analysis often requires measuring the fluorescence signal of GFP in a specific structure or area, to compare it with fluorescent intensities of other structures or areas. Once the quantities of fluorescent molecules in different sites or states are known, computer modelling can then be used to determine the parameter values (that is, the rate constants for binding interactions and exchange times) of the processes of interest¹⁰. Recent applications in which kinetic modelling has been used successfully include analysing the dynamics of nuclear proteins^{31,32,33,34,35}, protein transport through membrane trafficking pathways^27,36,37 and membrane coat protein dynamics³⁰.

Fig. 2
		Figure 2 | Photobleaching of GFP-tagged proteins to monitor dynamics.

Inverse FRAP. Inverse FRAP (iFRAP) is performed as a normal FRAP experiment with the exception that the molecules outside a region of interest are photobleached and the loss of fluorescence from the non-photobleached region is monitored over time. As opposed to the rate of recovery studied using a FRAP experiment, iFRAP offers a way to monitor the rate of movement out of a region. For example, iFRAP was used to monitor the dissociation kinetics of GFP-tagged RNA polymerase I components from sites of rRNA transcription³⁴. Because this method indirectly highlights a pool of molecules by decreasing the background fluorescence, it has also been used to follow Golgi to plasma membrane transport carriers as they moved from the Golgi and fused with the plasma membrane^27,37 (Fig. 2c).

Fluorescence localization after photobleaching. Fluorescence localization after photobleaching (FLAP)³⁸ also indirectly highlights a pool of molecules. For a FLAP experiment, the same protein-of-interest is tagged with two different fluorophores that co-localize when expressed in cells. By photobleaching one of these fluorophores, a selected pool can be highlighted and followed over time. Using cyan fluorescent protein (CFP)- and yellow fluorescent protein (YFP)-tagged -actin, monomer versus filamentous actin dynamics was demonstrated³⁸, and actin transport was monitored during cell protrusion³⁹. In another study, CFP- and YFP-tagged histone H2B molecules were co-expressed in cells undergoing mitosis⁴⁰. After one pool of the YFP-containing H2B molecules was photobleached, the movement of the non-photobleached pool was used to monitor chromosome positions throughout the cell cycle.

Fluorescence loss in photobleaching. Complementary to the photobleaching techniques discussed earlier, the continuity of a cell compartment can be monitored using a technique called fluorescence loss in photobleaching (FLIP) (Fig. 2b). In a FLIP experiment, a fluorescent cell is repeatedly photobleached within a small region while the whole cell is repeatedly imaged. Any regions of the cell that are connected to the area being bleached will gradually lose fluorescence due to lateral movement of mobile proteins into this area. By contrast, the fluorescence in unconnected regions will not be affected. In addition to assessing continuity between areas of the cell, FLIP can be used to assess whether a protein moves uniformly across a particular cell compartment or undergoes interactions that impede its motion^28,36,41. Furthermore, it can be used to reveal faint fluorescence in unconnected compartments that normally cannot be seen against the bright fluorescence that arises in other parts of the cell⁴².

Photobleaching applications

Photobleaching techniques that are applied to live-cell imaging are transforming our understanding of cellular organization and dynamics. For the first time, the mobility of diverse molecules in the cytoplasm, nucleus, organelle lumens and membranes of living cells can be measured, and the viscosity of these environments analysed. Moreover, resident components of organelles, once thought to be stable, have been shown to continuously enter and exit these structures. These findings are defining the biophysical characteristics of cellular compartments and their components, and are illuminating regulatory features of signalling and transport pathways.

Protein dynamics in the cytoplasm. The cytoplasm contains numerous macromolecular assemblies and cytoskeletal elements (including microtubules, actin and intermediate filaments). Yet, it has only recently become clear from FRAP studies that small molecules can rapidly diffuse through this system and bind reversibly to dynamic scaffolds. Such studies have shown that molecules up to 200 kDa undergo unhindered diffusion through the cytoplasm with D_eff values several times lower than those found in water^43,44. By contrast, larger molecules (200 kDa) or macromolecular complexes have impeded diffusion, presumably due to the extensive cytoskeletal meshwork of cells^44,45,46.

These diffusional properties have recently been shown to participate in the spatial organization and activity of signalling pathways. One example is the mitogen-activated protein kinase (MAPK) pathway. FRAP studies⁴⁷ that examined the dynamics of the MAPK Fus3 in yeast showed that it continuously binds to and dissociates from a plasma-membrane-localized scaffold molecule, Ste5. After being activated at the plasma membrane when bound to Ste5, Fus3 rapidly relocates to the nucleus by diffusion. The spatial localization of Fus3 activation and its dynamics at the plasma membrane are thought to help control and amplify MAPK signalling. A second example from yeast is the behaviour of septins — small GTPases that recruit proteins to form a ring at the cleavage site during cell division. Using FRAP, GFP-tagged septins in yeast were shown to be mobile during most of the cell cycle but then to become immobilized at the cleavage site at the time of budding⁴⁸. This leads to the recruitment of other proteins to this site, and thereby creates a diffusion barrier between mother and daughter cells⁴⁹. So, by changing between mobile and immobile states, septins help to control the temporal and spatial regulation of cytokinesis.

Protein dynamics in the nucleus. FRAP measurements of GFP-labelled nuclear proteins have revealed that many compartments in the nucleus — including nucleoli, Cajal bodies and splicing-factor compartments — are not stable entities but are steady-state assemblies of proteins that undergo continuous association and dissociation^{31,32,33,34,35}. The diffusion of proteins (including HMG17, SF2/ASF, fibrillarin, coilin and TBP) and the U7 small nuclear RNA (snRNA) in these subnuclear compartments are significantly lower (D_eff between 0.24–0.53 ΅m² sec^-1) than reported for freely diffusing peptides, GFP molecules or fluorescently labelled dextrans (>2 ΅m² sec^-1)^46,50. This indicates that exchange into and out of subnuclear compartments is the rate-limiting factor for the movement of these proteins and snRNA within the nucleus. In addition to providing insights into the dynamics of subnuclear structures, FRAP studies of the nucleus have revealed the kinetics of the binding of transcription-factor machinery to DNA promoters^51,52, the intranuclear mobility of messenger RNA⁵³ and the geography of chromosomes^40,54.

Intra- and inter-organelle dynamics. The micro-environment within organelles and the exchange of components between organelles have also been probed using FRAP. One example is the mitochondrial matrix, which has traditionally been thought of as too dense to allow the rapid movement of its components. However, FRAP measurements of the GFP-tagged matrix enzyme cytochrome oxidase c revealed that this small enzyme diffuses extremely rapidly in mitochondria⁵⁵. By contrast, components of the large macromolecular assemblies that comprise the fatty acid -oxidation pathway were immobilized, presumably through associations with the inner mitochondrial membrane⁵⁶. Based on these findings, it is thought that the clustered assemblies of proteins that are immobilized in the mitochondrial matrix provide a surface on which highly mobile substrates and enzymes can interact^55,56.

FRAP has also unveiled important characteristics of the ER lumen, which is enriched in molecules that are involved in protein biogenesis, folding and assembly. Under normal conditions, small soluble proteins can diffuse rapidly throughout the ER lumen with access to all areas^42,57. However, under conditions of cell stress — such as heat shock, change in osmolarity, calcium depletion, a glycosylation block or the production of unfolded proteins^42,58,59,60 — there are marked changes in the mobility of proteins and lumenal continuity. So, the ER lumen is not a stable environment, but undergoes significant global changes in response to cell stress, which could affect its numerous cellular roles.

FRAP techniques have been crucial for characterizing the mobility of GFP-tagged proteins that are embedded in organelle bilayers. The measured D_eff for many transmembrane proteins localized in the ER, Golgi apparatus or plasma membrane have values ranging from 0.2 to 0.5 ΅m² sec^-1 with little or no immobile fractions^41,42,61. This indicates that these proteins have unhindered lateral mobility in the membranes of these compartments. By contrast, large assemblies of membrane proteins in the ER (for example, translocons, TAP transporters and nuclear pores) or plasma-membrane proteins interacting with the extracellular matrix or cortical cytoskeletal elements, diffuse more slowly or have large immobile fractions^{25,60,62,63,64,65}. Studies of the diffusion properties of these molecules have important implications for understanding how proteins are retained in different membrane-bound compartments, and what mechanisms coordinate the processing and transport functions of membranes. For instance, alterations in TAP1–GFP D_eff under different peptide loads have provided evidence of TAP-complex conformational changes and interactions with class I major histocompatibility complex (MHC) molecules during peptide translocation^25,66. Additionally, FRAP experiments performed on the lamin-B receptor at various points during the cell cycle show that although the receptor is immobile in the nuclear envelope during interphase, it disperses to the endoplasmic reticulum (ER) and is completely mobile during mitosis⁶⁷.

Organelles of the secretory pathway, including the Golgi apparatus, have traditionally been thought to contain relatively stable resident components. But recent studies using FRAP techniques have revealed that membrane-bound and peripherally associated Golgi-resident proteins associate only transiently with this organelle⁶⁸. Whereas transmembrane enzymes can reside in the Golgi for up to 1–2 hours before recycling to the ER, peripherally associated coat and matrix proteins on the Golgi exchange with the soluble pools in the cytoplasm every 30–60 seconds. These results indicate that the Golgi apparatus is a highly dynamic organelle, the identity of which depends on continuous protein exchange with the cytoplasm and ongoing membrane input/output pathways.

Finally, photobleaching techniques have provided a powerful method for highlighting transport intermediates as they move along specific membrane-trafficking pathways^27,37,69,70 (Fig. 2b) and for analysing the dynamics of the protein-trafficking machinery^29,30,40,71. Studies of GFP-tagged components of membrane-trafficking machinery that sorts cargo into membrane-bound transport intermediates have shown that they undergo continuous binding to and dissociation from membranes irrespective of vesicle budding. These include COPII (Sec23/Sec24 and Sec13/ 31 heterodimers assembled onto ER membranes with the small GTPase, Sar1), COPI (a heptameric cytosolic protein complex recruited to Golgi membranes), Arf1 (a small GTPase) and clathrin (a major structural constituent forming the lattice around clathrin-coated vesicles). Whether this exchange represents a 'proof-reading' mechanism for ensuring proper loading of coated vesicles, or is necessary for lateral membrane differentiation into pleiomorphic transport intermediates⁷², remains to be investigated. In either case, the kinetics of this exchange have major implications for models of coat protein function and of the GTP binding and hydrolysis cycles of Arf1 and Sar1.

Photoactivation

Photoactivation is the photo-induced activation of an inert molecule to an active state. It is generally associated with the ultraviolet-light-induced release of a caging group from a 'caged' compound. Photoactivation of CAGED COMPOUNDS in the study of living cells is reviewed elsewhere⁷³, so our discussion is limited to recent advances in the development of genetically encoded photoactivatable fluorescent proteins.

Included in the development and discovery of new fluorescent protein variants^2,6,7 (see also the review on page S1 of this supplement) were attempts to produce photoactivatable fluorescent proteins. These studies yielded several molecules or techniques for optically highlighting proteins, but each had drawbacks for use in living cells, such as modest activation^74,75, low stability⁷⁶ or a requirement for low oxygen conditions^77,78. Recently, three photoactivatable fluorescent proteins — photoactivatable GFP (PA-GFP)⁷⁹, Kaede⁸⁰ and kindling fluorescent protein 1 (KFP1)⁸¹ — have been reported that offer improvements over the earlier versions.

The PA-GFP⁷⁹ was developed with the aim of optimizing the photoconversion properties of Aequorea victoria wtGFP⁷⁴, which produces only a threefold increase in fluorescence under 488 nm excitation. Mutation of threonine 203 to histidine in wtGFP to produce PA-GFP decreases the initial absorbance in the minor peak region (475 nm) and leads to 100-fold increase after photoactivation⁷⁹. Alternatively, for the Kaede protein, isolated from Trachyphyllia geoffroyi, photoactivation results in a 2,000-fold increase in its red-to-green fluorescence ratio⁸⁰. Finally, KFP1 — an A148G mutant (where A is alanine and G is glycine) of asFP595 (asCP, where 'FP' is fluorescent protein and 'CP' is chromoprotein) from the sea anemone, Anemonia sulcata — gives a 30-fold increase in red fluorescence after photoactivation⁸¹.

All of these molecules share the common characteristic of displaying low levels of fluorescence before photoactivation and higher levels after photoactivation. In a typical experiment, a cell or organism that is expressing the photoactivatable fluorescent protein is imaged at one wavelength prior to, and at various intervals after, photoactivation of a selected region with a different wavelength. However, the properties of each protein, including the wavelengths used for imaging and photoactivation, offer distinct advantages and disadvantages. For example, PA-GFP and Kaede both require 400 nm light for photoactivation, whereas KFP1 uses green light (532 nm), which is probably better for use with living cells. Kaede displays a remarkable 2,000-fold increase in its red-to-green fluorescence ratio, but the use of both the red and green fluorescence bands could make multilabel experiments problematic. On the other hand, the green fluorescence of Kaede is bright enough to visualize the localization of the non-photoactivated proteins easily, whereas visualizing PA-GFP and KFP1 is more problematic due to their low fluorescence before photoactivation. The self-association properties of Kaede and KFP1 into tetrameric forms limit their usefulness as protein fusion tags because tetramerization might perturb parent protein localization and trafficking. The recent engineering of the DsRed protein into a monomeric form⁸² is encouraging for the possibility of the eventual disruption of Kaede and KFP1 into monomers. Variants that are derived from A. victoria, such as PA-GFP, self-associate to a lesser degree, and even those interactions can be disrupted by one of three further point mutations⁸³. Because of this, PA-GFP can be used as a reliable tag for creating fluorescent reporter molecules.

The ability to 'switch on' the fluorescence of the photoactivatable proteins makes them excellent tools for exploring protein behaviour in living cells. As the fluorescence of these proteins comes only after photoactivation, newly synthesized non-photoactivated pools are unobserved and do not complicate experimental results (Fig. 3a). This signal independence from new protein synthesis could allow the study of protein degradation of tagged molecules by 'optical pulse labelling' and monitoring of the fluorescence over time (Fig. 3b). Photoactivation of these proteins is generally rapid and gives stable fluorescence signals. Therefore, they can be used to examine various kinetic properties of tagged proteins, such as their D_eff , M_f , compartmental residency time and exchange. Lastly, cell lineage or movement in a developing organism can be monitored by imaging the fluorescence dispersion after photoactivation of a single cell or subpopulation of cells⁸¹ (Fig. 3c). So, these proteins have remarkable promise to complement and extend the range of present fluorescent-protein imaging applications.

Fig. 3
		Figure 3 | Photoactivation of fluorescent proteins.

Concluding remarks

The battery of fluorescent proteins and imaging tools that allow us to monitor protein dynamics in living cells continue to provide numerous new insights into the behaviour of proteins, organelles and cells. In so doing, they have ushered in a new era of cell biology in which kinetic microscopy methods can be used to decipher pathways and mechanisms of biological processes. The microscopy techniques of photobleaching and photoactivation are perhaps the most versatile and widely used of these methods. Their ability to alter the fluorescence steady state without perturbing protein dynamics offers unprecedented opportunities for obtaining quantitative information about protein concentrations, diffusion rates, binding kinetics and protein lifetimes in single live cells, which have been indiscernible using traditional biochemical approaches. Such information is paramount to understanding how biological processes unfold, are regulated and interact in the living cell.

Looking to the future, photobleaching and photoactivation will almost certainly continue to provide important new results as their applications are extended by the development of newer instruments that push the limits of temporal and spatial detection, and by the discovery of brighter and differently coloured fluorescent proteins. For example, FRAP can be combined with other microscopic imaging approaches, including two-photon microscopy⁸⁴, (see MULTI-PHOTON MICROSCOPY) or TOTAL INTERNAL REFLECTION microscopy⁸⁵, to study events at specific sites in the cell. And, photobleaching or photoactivation can be combined with fluorescence energy-transfer techniques^2,8 to study protein interactions with greater precision. These advances will continue to require computational approaches to comprehend the plethora of quantitative experimental data¹⁰, as well as new database tools for the analysis of specific models and their relationship to other more complex models.

References

1.	van Roessel, P. & Brand, A. H. Imaging into the future: visualizing gene expression and protein interactions with fluorescent proteins. Nature Cell Biol. 4, E15–E20 ( 2002). | Article | | PubMed |
2.	Zhang, J., Campbell, R. E., Ting, A. Y. & Tsien, R. Y. Creating new fluorescent probes for cell biology. Nature Rev. Mol. Cell Biol. 3, 906–918 ( 2002). | Article | | PubMed |
3.	Chalfie, M., Tu, Y., Euskirchen, G., Ward, W. W. & Prasher, D. C. Green fluorescent protein as a marker for gene expression. Science 263, 802–805 (1994). | PubMed |
4.	Gonzαlez, C. & Bejarano, L. A. Protein traps: using intracellular localization for cloning. Trends Cell Biol. 10, 162–165 (2000). | Article | | PubMed |
5.	Lippincott-Schwartz, J., Roberts, T. H. & Hirschberg, K. Secretory protein trafficking and organelle dynamics in living cells. Annu. Rev. Cell Dev. Biol. 16, 557–589 (2000). | Article | | PubMed |
6.	Tsien, R. Y. The green fluorescent protein. Annu. Rev. Biochem. 67, 509–544 (1998). | Article | | PubMed |
7.	Lippincott-Schwartz, J. & Patterson, G. H. Development and use of fluorescent protein markers in living cells. Science 300, 87–91 (2003). | Article | | PubMed |
8.	Lippincott-Schwartz, J., Snapp, E. & Kenworthy, A. Studying protein dynamics in living cells. Nature Rev. Mol. Cell Biol. 2, 444–456 ( 2001). | Article | | PubMed |
9.	Houtsmuller, A. B. & Vermeulen, W. Macromolecular dynamics in living cell nuclei revealed by fluorescence redistribution after photobleaching. Histochem. Cell Biol. 115, 13–21 ( 2001).
10.	Phair, R. D. & Misteli, T. Kinetic modeling approaches to in vivo imaging. Nature Rev. Mol. Cell Biol. 2, 898–907 ( 2001). | Article |
11.	Klonis, N. et al. Fluorescence photobleaching analysis for the study of cellular dynamics. Eur. Biophys. J. 31, 36–51 ( 2002). | Article |
12.	Axelrod, D., Koppel, D. E., Schlessinger, J., Elson, E. & Webb, W. W. Mobility measurement by analysis of fluorescence photobleaching recovery kinetics. Biophys. J. 16, 1055–1069 ( 1976). | PubMed |
13.	Elson, E. L., Schlessinger, J., Koppel, D. E., Axelrod, D. & Webb, W. W. Measurement of lateral transport on cell surfaces. Prog. Clin. Biol. Res. 9, 137–147 ( 1976).
14.	Jacobson, K., Derzko, Z., Wu, E. S., Hou, Y. & Poste, G. Measurement of the lateral mobility of cell surface components in single, living cells by fluorescence recovery after photobleaching. J. Supramol. Struct. 5, 565(417)–576(428) ( 1976).
15.	Schlessinger, J. et al. Lateral transport on cell membranes: mobility of concanavalin A receptors on myoblasts. Proc. Natl Acad. Sci. USA 73, 2409–2013 ( 1976). | PubMed |
16.	Schindler, M., Osborn, M. J. & Koppel, D. E. Lateral diffusion of lipopolysaccharide in the outer membrane of Salmonella typhimurium . Nature 285, 261–263 ( 1980).
17.	Sheetz, M. P., Schindler, M. & Koppel, D. E. Lateral mobility of integral membrane proteins is increased in spherocytic erythrocytes. Nature 285, 510–511 ( 1980).
18.	Reits, E. A. J. & Neefjes, J. J. From fixed to FRAP: measuring protein mobility and activity in living cells. Nature Cell Biol. 3, E145–E147 ( 2001). | Article | | PubMed |
19.	Kao, H. P., Abney, J. R. & Verkman, A. S. Determinants of the translational mobility of a small solute in cell cytoplasm. J. Cell Biol. 120, 175–184 ( 1993). | PubMed |
20.	Gordon, G. W., Chazotte, B., Wang, X. F. & Herman, B. Analysis of simulated and experimental fluorescence recovery after photobleaching. Data for two diffusing components. Biophys. J. 68, 766–778 ( 1995). | PubMed |
21.	Verkman, A. S. Solute and macromolecule diffusion in cellular aqueous compartments. Trends Biochem. Sci. 2, 27–33 ( 2002). | Article |
22.	Siggia, E. D., Lippincott-Schwartz, J. & Bekiranov, S. Diffusion in inhomogeneous media: theory and simulations applied to whole cell photobleach recovery. Biophys. J. 79, 1761–1770 ( 2000). | PubMed |
23.	Snapp, E. L., Altan, N. & Lippincott-Schwartz, J. in Current Protocols in Cell Biology (eds Bonifacino, J., Dasso, M., Harford, J. B., Lippincott-Schwartz, J. & Yamada, K. M.) 21.1.1–21.1.23 (John Wiley & Sons, Inc., New York, 2003).
24.	Periasamy, N. & Verkman, A. S. Analysis of fluorophore diffusion by continuous distributions of diffusion coefficients: application to photobleaching measurements of multicomponent and anomalous diffusion. Biophys. J. 75, 557–567 ( 1998). | PubMed |
25.	Marguet, D. et al. Lateral diffusion of GFP-tagged H2Ld molecules and of GFP-TAP1 reports on the assembly and retention of these molecules in the endoplasmic reticulum. Immunity 11, 231–240 ( 1999). | PubMed |
26.	Wu, E. S., Jacobson, K., Szoka, F. & Portis, J. A. Lateral diffusion of a hydrophobic peptide, N-4-nitrobenz-2-oxa-1,3-diazole gramicidin S, in phospholipid multibilayers. Biochemistry 17, 5543–5550 ( 1978).
27.	Hirschberg, K. et al. Kinetic analysis of secretory protein traffic and characterization of Golgi to plasma membrane transport intermediates in living cells. J. Cell Biol. 143, 1485–1503 ( 1998). | Article | | PubMed |
28.	Phair, R. D. & Misteli, T. High mobility of proteins in the mammalian nucleus. Nature 404, 604–609 ( 2000). | Article | | PubMed |
29.	Wu, X. et al. Clathrin exchange during clathrin-mediated endocytosis. J. Cell Biol. 155, 291–300 ( 2001). | Article | | PubMed |
30.	Presley, J. F. et al. Dissection of COPI and Arf1 dynamics in vivo and role in Golgi membrane transport. Nature 417, 187–193 ( 2002). | Article | | PubMed |
31.	Misteli, T., Caceres, J. F. & Spector, D. L. The dynamics of a pre-mRNA splicing factor in living cells. Nature 387, 523–527 ( 1997). | Article | | PubMed |
32.	Snaar, S., Wiesmeijer, K., Jochemsen, A. G., Tanke, H. J. & Dirks, R. W. Mutational analysis of fibrillarin and its mobility in living human cells. J. Cell Biol. 151, 653–662 ( 2000). | Article | | PubMed |
33.	Chen, D., Hinkley, C. S., Henry, R. W. & Huang, S. TBP dynamics in living human cells: constitutive association of TBP with mitotic chromosomes. Mol. Biol. Cell 13, 276–284 ( 2002). | Article | | PubMed |
34.	Dundr, M. et al. A kinetic framework for a mammalian RNA polymerase in vivo . Science 298, 1623–1626 ( 2002). | Article |
35.	Handwerger, K. E., Murphy, C. & Gall, J. G. Steady-state dynamics of Cajal body components in the Xenopus germinal vesicle. J. Cell Biol. 160, 495–504 (2003). | Article |
36.	Zaal, K. J. M. et al. Golgi membranes are absorbed into and reemerge from the ER during mitosis. Cell 99, 589–601 (1999). | PubMed |
37.	Nichols, B. J. et al. Rapid cycling of lipid raft markers between the cell surface and Golgi complex. J. Cell Biol. 153, 529–541 (2001). | Article | | PubMed |
38.	Dunn, G. A., Dobbie, I. M., Monypenny, J., Holt, M. R. & Zicha, D. Fluorescence localization after photobleaching (FLAP): a new method for studying protein dynamics in living cells. J. Microsc. 205, 109–112 (2002). | Article | | PubMed |
39.	Zicha, D. et al. Rapid actin transport during cell protrusion. Science 300, 142–145 (2003). | Article |
40.	Gerlich, D. et al. Global chromosome positions are transmitted through mitosis in mammalian cells. Cell 112, 751–764 (2003). | PubMed |
41.	Cole, N. B. et al. Diffusional mobility of Golgi proteins in membranes of living cells. Science 273, 797–801 ( 1996). | PubMed |
42.	Nehls, S. et al. Dynamics and retention of misfolded proteins in native ER membranes. Nature Cell Biol. 2, 288–295 ( 2000). | Article | | PubMed |
43.	Luby-Phelps, K., Taylor, D. L. & Lanni, F. Probing the structure of cytoplasm. J. Cell Biol. 102, 2015–2022 ( 1986).
44.	Swaminathan, R., Hoang, C. P. & Verkman, A. S. Photobleaching recovery and anisotropy decay of green fluorescent protein GFP-S65T in solution and cells: cytoplasmic viscosity probed by green fluorescent protein translational and rotational diffusion. Biophys. J. 72, 1900–1907 ( 1997). | PubMed |
45.	Luby-Phelps, K., Castle, P. E., Taylor, D. L. & Lanni, F. Hindered diffusion of inert tracer particles in the cytoplasm of mouse 3T3 cells. Proc. Natl Acad. Sci. USA 84, 4910–4913 ( 1987). | PubMed |
46.	Seksek, O., Biwersi, J. & Verkman, A. S. Translational diffusion of macromolecule-sized solutes in cytoplasm and nucleus. J. Cell Biol. 138, 131–142 ( 1997). | Article | | PubMed |
47.	van Drogen, F., Stucke, V. M., Jorritsma, G. & Peter, M. MAP kinase dynamics in response to pheromones in budding yeast. Nature Cell Biol. 3, 1051–1059 ( 2001). | Article |
48.	Dobbelaere, J., Gentry, M. S., Hallberg, R. L. & Barral, Y. Phosphorylation-dependent regulation of septin dynamics during the cell cycle. Dev. Cell 4, 345–357 ( 2003).
49.	Faty, M., Fink, M. & Barral, Y. Septins: a ring to part mother and daughter. Curr. Genet. 41, 123–131 ( 2002). | Article |
50.	Reits, E. et al. Peptide diffusion, protection, and degradation in nuclear and cytoplasmic compartments before antigen presentation by MHC Class I. Immunity 18, 97–108 ( 2003). | PubMed |
51.	Becker, M. et al. Dynamic behavior of transcription factors on a natural promoter in living cells. EMBO Rep. 3, 1188–1194 ( 2002). | Article |
52.	Kimura, H., Sugaya, K. & Cook, P. R. The transcription cycle of RNA polymerase II in living cells. J. Cell Biol. 159, 777–782 ( 2002). | Article | | PubMed |
53.	Calapez, A. et al. The intranuclear mobility of messenger RNA binding proteins is ATP dependent and temperature sensitive. J. Cell Biol. 159, 795–805 ( 2002). | Article |
54.	Walter, J., Schermelleh, L., Cremer, M., Tashiro, S. & Cremer, T. Chromosome order in HeLa cells changes during mitosis and early G1, but is stably maintained during subsequent interphase stages. J. Cell Biol. 160, 685–697 ( 2003). | Article | | PubMed |
55.	Partikian, A., Φlveczky, B., Swaminathan, R., Li, Y. & Verkman, A. S. Rapid diffusion of green fluorescent protein in the mitochondrial matrix. J. Cell Biol. 140, 821–829 ( 1998). | Article | | PubMed |
56.	Haggie, P. M. & Verkman, A. S. Diffusion of tricarboxylic acid cycle enzymes in the mitochondrial matrix in vivo. Evidence for restricted mobility of a multienzyme complex. J. Biol. Chem. 277, 40782–40788 ( 2002). | Article |
57.	Dayel, M. J., Hom, E. F. Y. & Verkman, A. S. Diffusion of green fluorescent protein in the aequous-phase lumen of endoplasmic reticulum. Biophys. J. 76, 2843–2851 ( 1999). | PubMed |
58.	Subramanian, K. & Meyer, T. Calcium-induced restructuring of nuclear envelope and endoplasmic reticulum calcium stores. Cell 89, 963–971 ( 1997). | PubMed |
59.	Terasaki, M. Dynamics of the endoplasmic reticulum and Golgi apparatus during early sea urchin development. Mol. Biol. Cell 11, 897–914 ( 2000). | PubMed |
60.	Nikonov, A. V., Snapp, E. L., Lippincott-Schwartz, J. & Kreibich, G. Active translocon complexes labeled with GFP-Dad1 diffuse slowly as large polysome arrays in the endoplasmic reticulum. J. Cell Biol. 158, 497–506 ( 2002). | Article |
61.	Schmoranzer, J., Goulian, M., Axelrod, D. & Simon, S. M. Imaging constitutive exocytosis with total internal reflection fluorescence microscopy. J. Cell Biol. 149, 23–32 ( 2000). | Article | | PubMed |
62.	Barbour, S. & Edidin, M. Cell-specific constraints to the lateral diffusion of a membrane glycoprotein. J. Cell Physiol. 150, 526–533 ( 1992).
63.	Chakrabarti, A., Matko, J., Rahman, N. A., Barisas, B. G. & Edidin, M. Self-association of class I major histocompatibility complex molecules in liposome and cell surface membranes. Biochemistry 31, 7182–7189 ( 1992).
64.	Daigle, N. et al. Nuclear pore complexes form immobile networks and have a very low turnover in live mammalian cells. J. Cell Biol. 154, 71–84 ( 2001). | Article | | PubMed |
65.	Griffis, E. R., Altan, N., Lippincott-Schwartz, J. & Powers, M. A. Nup98 is a mobile nucleoporin with transcription-dependent dynamics. Mol. Biol. Cell 13, 1282–1297 ( 2002). | Article | | PubMed |
66.	Reits, E. A. J., Vos, J. C., Grommι, M. & Neefjes, J. The major substrates for TAP in vivo are derived from newly synthesized proteins. Nature 404, 774–778 ( 2000). | Article | | PubMed |
67.	Ellenberg, J. et al. Nuclear membrane dynamics and reassembly in living cells: targeting of an inner nuclear membrane protein in interphase and mitosis. J. Cell Biol. 138, 1193–1206 ( 1997). | Article | | PubMed |
68.	Ward, T. H., Polishchuk, R. S., Caplan, S., Hirschberg, K. & Lippincott-Schwartz, J. Maintenance of Golgi structure and function depends on the integrity of ER export. J. Cell Biol. 155, 557–570 ( 2001). | Article | | PubMed |
69.	Presley, J. F. et al. ER-to-Golgi transport visualized in living cells. Nature 389, 81–85 (1997). | Article | | PubMed |
70.	Nakata, T., Terada, S. & Hirokawa, N. Visualization of the dynamics of synaptic vesicle and plasma membrane proteins in living axons. J. Cell Biol. 140, 659–674 ( 1998). | Article | | PubMed |
71.	Stephens, D. J., Lin-Marq, N., Pagano, A., Pepperkok, R. & Paccaud, J. P. COPI-coated ER-to-Golgi transport complexes segregate from COPII in close proximity to ER exit sites. J. Cell Sci. 113, 2177–2185 ( 2000). | PubMed |
72.	Bonifacino, J. & Lippincott-Schwartz, J. Coat proteins: shaping membrane transport. Nature Rev. Mol. Cell Biol. 4, 409–414 ( 2003). | Article |
73.	Politz, J. C. Use of caged fluorophores to track macromolecular movement in living cells. Trends Cell Biol. 9, 284–287 ( 1999). | Article | | PubMed |
74.	Yokoe, H. & Meyer, T. Spatial dynamics of GFP-tagged proteins investigated by local fluorescence enhancement. Nature Biotechnol. 14, 1252–1256 ( 1996). | PubMed |
75.	Marchant, J. S., Stutzmann, G. E., Leissring, M. A., LaFerla, F. M. & Parker, I. Multiphoton-evoked color change of DsRed as an optical highlighter for cellular and subcellular labeling. Nature Biotechnol. 19, 645–649 ( 2001). | Article | | PubMed |
76.	Lukyanov, K. A. et al. Natural animal coloration can be determined by a nonfluorescent green fluorescent protein homolog. J. Biol. Chem. 275, 25879–25882 ( 2000). | Article | | PubMed |
77.	Elowitz, M. B., Surette, M. G., Wolf, P. -E., Stock, J. & Leibler, S. Photoactivation turns green fluorescent protein red. Curr. Biol. 7, 809–812 ( 1997). | PubMed |
78.	Sawin, K. E. & Nurse, P. Photoactivation of green fluorescent protein. Curr. Biol. 7, R606–R607 ( 1997). | PubMed |
79.	Patterson, G. H. & Lippincott-Schwartz, J. A photoactivatable GFP for selective photolabeling of proteins and cells. Science 297, 1873–1877 ( 2002). | Article | | PubMed |
80.	Ando, R., Hama, H., Yamamoto-Hino, M., Mizuno, H. & Miyawaki, A. An optical marker based on the UV-induced green-to-red photoconversion of a fluorescent protein. Proc. Natl Acad. Sci. USA 99, 12651–12656 ( 2002). | Article | | PubMed |
81.	Chudakov, D. M. et al. Kindling fluorescent proteins for precise in vivo photolabeling. Nature Biotechnol. 21, 191–194 ( 2003). | Article |
82.	Campbell, R. E. et al. A monomeric red fluorescent protein. Proc. Natl Acad. Sci. USA 99, 7877–7882 ( 2002). | Article | | PubMed |
83.	Zacharias, D. A., Violin, J. D., Newton, A. C. & Tsien, R. Partitioning of lipid-modified monomeric GFPs into membrane microdomains of live cells. Science 296, 913–916 ( 2002). | Article | | PubMed |
84.	Denk, W., Piston, D. W. & Webb, W. W. in Handbook of Biological Confocal Microscopy (ed. Pawley, J. B.) 445–458 (Plenum Press, New York, 1995).
85.	Steyer, J. A. & Almers, W. A real-time view of life within 100 nm of the plasma membrane. Nature Rev. Mol. Cell Biol. 2, 268–275 ( 2001). | Article | | PubMed |
86.	Levin, M. H., Haggie, P. M., Vetrivel, L. & Verkman, A. S. Diffusion in the endoplasmic reticulum of an aquaporin-2 mutant causing human nephrogenic diabetes insipidus. J. Biol. Chem. 276, 21331–21336 ( 2001). | Article | | PubMed |
87.	Lippincott–Schwartz, J. et al. in Green Fluorescent Proteins (eds Sullivan, K. F. & Kay, S. A.) 261–281 (Academic Press, San Diego, 1999).