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ScienceDirect Live-cell reporters for fluorescence imaging Ivan R Correˆa Jr Advances in the development of new fluorescent reporters and imaging techniques have revolutionized our ability to directly visualize biological processes in living systems. Real-time analysis of protein localization, dynamics, and interactions has been made possible by site-specific protein labeling with custom designed probes. This review outlines some of the most recent advances in the design and application of live-cell imaging probes, with a particular focus on SNAP-tag technology. Specific examples illustrating applications in superresolution and single-molecule imaging, protein trafficking and recycling, and protein–protein interactions are presented. Addresses New England Biolabs, Inc., 240 County Road, Ipswich, MA 01938, USA Corresponding author: Correˆa, Ivan R ([email protected])

Current Opinion in Chemical Biology 2014, 20:36–45 This review comes from a themed issue on Molecular imaging Edited by Christian Eggeling and Mike Heilemann

http://dx.doi.org/10.1016/j.cbpa.2014.04.007 1367-5931/# 2014 Elsevier Ltd. All rights reserved.

Introduction Fluorescence microscopy is a powerful tool for studying the dynamics and localization of proteins in living cells. Proteins are routinely imaged by fusion to auto-fluorescent proteins or to protein/peptide tags that can be site-specifically labeled with synthetic fluorophores. The success of any strategy for live-cell imaging lies in the ability to specifically confer the desired optical properties to the protein of interest, thereby providing means to visualize and interrogate the protein in its native environment. Tagbased approaches utilize specific recognition sequences to recruit chemical probes for in situ labeling of the target protein. The binding of the probe to the protein tag occurs either through a tag-mediated self-labeling reaction or is assisted by an auxiliary enzyme. A key advantage of sitespecific labeling over imaging approaches on the basis of auto-fluorescence proteins is the ability to use chemistry to modulate the biophysical properties of a given synthetic fluorophore to the needs and constraints imposed by the experiment. Moreover, the high spatial and temporal resolution, molecular specificity, and nondestructive compatibility with living systems make site-specific labeling Current Opinion in Chemical Biology 2014, 20:36–45

suitable for a broad range of applications from in vivo imaging to drug discovery processes. Various peptide and protein fusion tags have been developed that permit the study of proteins in live cells and organisms, including self-labeling systems such as Tetracysteine-tag, SNAP-tag [1,2], CLIP-tag [3], HaloTag [4], TMP-tag [5] and BL-tag [6], and the enzyme-mediated systems such as the ones on the basis of phosphopantetheinyl transferases (AcpS and Sfp) [7], sortase (SrtA) [8], and lipoic acid ligase (LplA) [9]. In this review, I will focus on the development and recent applications of live-cell fluorescence imaging reporters for one particular site-specific labeling technique, SNAP-tag. For further information on other site-specific labeling approaches as well as on the use of auto-fluorescent proteins, the reader is encouraged to consult some excellent reviews published in recent years [10,11,12]. The SNAP-tag is an engineered mutant of the human repair protein O6-alkylguanine-DNA alkyltransferase (hAGT) which reacts with O6-benzylguanine (BG) [1,2] or O6-benzyl-4-chloropyrimidine (CP) [13] substrates modified with a synthetic label (e.g., fluorophores, quantum dots, affinity ligands, and gold nanoparticles). The label is covalently attached to the fusion tag in a welldefined mechanism, predictable stoichiometry, and rapid kinetics (Figure 1a) [14]. Selective labeling of intracellular or membrane proteins can be achieved by the appropriate selection of cell-permeable or cell-impermeable substrates, respectively. The SNAP-tag protein labeling system is distinguished from auto-fluorescence proteinbased approaches by the wealth of different reporter probes it makes available for imaging (e.g., fluorophores, quantum dots, affinity ligands, gold nanoparticles, etc.) as well as the easy synthesis of custom probes for new applications, such as fluorogenic probes [15,16], photoactivatable fluorophores [17–19], as well as fluorescent sensors for metal ions [20,21] and cell metabolites [22– 24]. Additionally, the ability to noninvasively label proteins in any compartment of the cell, including the nucleus, cytoplasm, and at both faces of the plasma membrane, significantly expands the applications into a wide variety of experimental settings in cell biology [25]. SNAP-tag has been successfully utilized in many cellular and in vivo imaging studies, including analysis of protein function [26], protein half-life [27], protein trafficking and recycling [28], protein–protein interactions [29], and protein–drug interactions [30,31]. Its high labeling specificity allied with the superior optical properties of synthetic fluorophores has greatly increased our ability to study proteins in living systems. Here I will discuss the design of new fluorescent reporters and present examples of recent applications, such as those for pulse-chase www.sciencedirect.com

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experiments, multi-color protein imaging, superresolution microscopy, and tracking of single protein molecules in live-cell experiments.

Design of new live-cell reporters for fluorescence imaging Fluorogenic reporters

Fluorogenic reporters allow for direct monitoring of fluorescence signals with high sensitivity and spatial resolution by minimizing the fluorescence background caused by unreacted or nonspecifically bound probes. Fluorogenic (‘dark’ or ‘quenched’) probes exhibit nearly no basal fluorescence, but generate an intense fluorescence response after they have bound to their intended target, thereby resulting in much higher signal-to-noise ratios than conventional fluorophores. We and others have developed SNAP-tag fluorogenic substrates to enable real-time imaging of dynamic cellular processes, such as protein expression, localization, trafficking and degradation [15,16]. These substrates consist of a benzylguanine scaffold bearing a fluorophore attached at the para position of the benzylic ring and a dark quencher attached at the C-8 position of the guanine ring (this position was shown to be the only one amenable for chemical modification without significant deterioration of probe reactivity towards SNAP-tag) (Figure 1b). The fluorescence emission of the reporter fluorophore is intramolecularly quenched through a combination of Fo¨rster-type resonance energy transfer (FRET) and photo-induced electron transfer (PET) [16]. Upon reaction with the tagged protein, the quencher group is dissociated leading to up to 50-fold increase in the relative fluorescence intensity of the fluorophore. By turning ‘on’ the fluorescence signal only at the target protein, fluorogenic reporters meet the critical need for systematic ‘no-wash’ before fluorescence imaging measurements (Figure 1c). The usefulness of fluorogenic reporters to continually monitor the spatiotemporal dynamics of the EGF receptor (EGFR) during cell migration, without the need for washing the cells, was demonstrated by Urano and coworkers [15]. Of particular interest is the development of near-infrared fluorogenic reporters, which could virtually eliminate background fluorescence in live-animal or deep tissue analysis, and therefore, circumvent probe clearance issues for real-time imaging. In attempt to achieve this goal, SNAP-tag fluorogenic substrates carrying a near-infrared emitting fluorophore (IRDye 800CW) and a non-fluorescent broad range quencher (IRDye QC-1) were synthesized; however, these probes failed to detect xenograph tumors expressing SNAP-tag fused to the ADRb2 adrenergic receptor, presumably due to the reduced reaction rate of these substrates [32]. More recently, Johnsson and co-workers have introduced a novel class of cell-permeable near-infrared fluorescent reporters on the basis of a silicon–rhodamine (SiR) fluorwww.sciencedirect.com

ophore which permits the imaging of proteins in living cells without washing steps [33]. The fluorogenic character of the probe stems from an equilibrium between its fluorescent zwitterionic and non-fluorescent spirolactone forms (Figure 2a). Coupling of the SiR probe to the protein tag favors the formation of the fluorescent zwitterion, whereas the aggregation of unreacted probe and nonspecific binding to hydrophobic structures favors the formation of the non-fluorescent spirolactone, thereby resulting in minimal background staining in live-cell experiments. The high sensitivity of the SiR fluorophore at a wavelength that exhibits low cellular autofluorescence and phototoxicity has enabled the labeling of SNAP-tag in cortical neurons in rat-brain sections. Furthermore, the excellent brightness and photostability of this probe were confirmed in long-term imaging experiment in which cells that expressed SNAP-tag fusions were grown in the presence of the probe and imaged continuously over 48 h. The utility of SiR for live-cell superresolution microscopy has also been demonstrated [33].

Photoactivatable fluorescent reporters Dynamic measurements often require spatial tracking of a molecule or an entire organelle within a cell. The tools of choice for these purposes are photoactivatable or photoswitchable fluorescent reporters. These probes are sensitive to photoinduced conversions, such that their optical properties are modified upon irradiation with light of a specific wavelength. Photoactivatable probes show fluorescence quantum yield enhancement, whereas photoswitchable probes exhibit emission wavelength switching. The possibility to actively control the fluorescence emission at a time in a diffraction-limited region allow for the acquisition of superresolution images for nanoscale visualization. Photoinduced fluorescent proteins have often been used for live-cell superresolution microscopy because of their ability of being genetically targeted; however, they provide 10-fold fewer photons before photobleaching than good small-molecule emitters [34]. Single-molecule-based imaging by means of photoactivatable fluorescent proteins, particularly in the green region of the optical spectrum, suffers from high background and low contrast ratios. As sitespecific labeling technologies, such as the SNAP-tag, allow the spatial targeting of synthetic fluorophores to specific subcellular locations, they can be advantageously employed for visualizing, monitoring, and quantifying dynamic molecular events in living cells with high spatial and temporal resolution. SNAP-tag photoinduced substrates have been generated by a so-called ‘caging’ process in which a fluorophore of choice is converted into a non-fluorescent state by the incorporation of a photocleavable group, for example 4,5dimethoxy-2-nitrobenzyl (DMNB) (Figure 2b) [35]. Fluorophore uncaging and consequent fluorescence Current Opinion in Chemical Biology 2014, 20:36–45

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(a) Schematic representation of the labeling reaction of SNAP-tag with fluorescent probes. SNAP-tag reacts with O6-benzylguanine (BG) substrates resulting in the covalent attachment of a fluorescent label to the active site cysteine. (b) Labeling of SNAP-tag fusion proteins with fluorogenic probes.

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Examples of fluorescent reporters for labeling and imaging SNAP-tag fusion proteins. (a) A SNAP-tag near-infrared silicon–rhodamine (SiR) substrate. Structures of its fluorescent zwitterionic and non-fluorescent spirolactone forms. (b) Caged rhodamine BG-cRhod. A photocleavable 4,5-dimethoxy-2nitrobenzyl group (DMNB) is attached to the fluorophore core via a carbamate bond forcing the probe in its non-fluorescent lactone configuration. The fluorescence is restored through irradiation with UV light. (c) A cleavable SNAP-tag fluorescent substrate. After labeling a SNAP-tag fusion protein with a cleavable reporter group, the fluorescent label can be readily released by reduction of the disulfide bond with tris(2-carboxyethyl)phosphine (TCEP) or sodium 2-sulfanylethanesulfonate (MESNA). (d) A SNAP-tag bifunctional reporter comprised of a biotin moiety and a fluorescent dye designed for concurrent imaging and immobilization/affinity purification protocols.

recovery can be achieved with high spatial and temporal resolution through irradiation with UV light. The increase in fluorescence intensity upon uncaging SNAP-tag bound to rhodamine and BODIPY caged fluorophores was of 200-fold and 600-fold, respectively. Caged fluorophores such as these have become important tools in investigating dynamic biological processes, including tracking of cell lineage in zebrafish and protein trafficking [18,36]. Another attractive application of caged fluorophores is for imaging of cellular structures with nanometer resolution by means of photoactivated localization microscopy (PALM) [35]. A critical limitation of the available caged probes is that they can only be used to label proteins on the surface of living cells. Labeling of intracellular targets currently requires prior fixation/permeabilization of cells or the use of invasive cell-loading techniques, such as bead-loading [19], microinjection [18,37] or electroporation [38].

Cleavable fluorescent reporters Membrane receptor internalization, sorting, and recycling are essential to the proper regulation of cell signaling pathways. Biochemical approaches, in particular receptor biotinylation and immunofluorescence labeling methods, have played a critical role in elucidating some of the mechanisms underlying receptor trafficking. These approaches will continue to be important for microscopic assessment of membrane receptors; however, new imaging tools are needed to facilitate real-time visualization of the subcellular organization and dynamics of receptormediated signaling systems. To this end, SNAP-tag fluorescent reporters incorporating a reducible disulfide bond between the benzylguanine group and the fluorophore of choice (Figure 2c) have been designed to permit the study of trafficking and recycling events in living cells [39]. These probes allow the user to chemically release a fluorophore from the labeled SNAP-tag fusion. In a typical

(Figure 1 Legend Continued) A quencher attached at the guanine ring is released in the course of the labeling reaction. (c) Imaging with the SNAP-tag technology. First, the protein of interest is subcloned and expressed as a SNAP-tag fusion. The tagged protein is then labeled with a fluorescent probe. Cells are extensively washed to remove excess probe before imaging. Fluorogenic probes can be directly imaged without the need for washing steps. www.sciencedirect.com

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experiment, cell surface receptors are labeled with a cleavable fluorescent probe and internalized in the presence of an appropriate agonist. Subsequent incubation with a cell-impermeable reducing agent (such as TCEP or MESNA) removes any remaining cell surface-associated fluorescence due to non-internalized receptors without affecting the population of internalized labeled receptors. This significantly reduces the background fluorescence at the surface of living cells, making the approach particularly useful for studying receptor sorting and recycling dynamics within endosomes and the plasma membrane. Donaldson and co-workers implemented this approach for tracking the intracellular fate of internalized G protein-coupled receptors (GPCRs), including the b2 adrenergic receptor (b2ADR) and neurokinin-1 receptor (NK1R) [39], and the human interleukin-2 receptor [40].

Multifunctional reporters Multifunctional reporters can contain either multiple reactive groups (which are specific for each self-labeling technology) or multiple labels (fluorophores, affinity ligands, etc.) in the same molecule. Reporter probes comprised of multiple reactive groups have been devised to crosslink proteins as an alternative to FRET approaches aimed at the investigation of protein–protein interactions in living cells and in cell extracts. These probes can be homobifunctional, for the study of identical protein monomers, or heterobifunctional, for the dimerization processes between different proteins. The reactive groups in a crosslinking substrate are usually joined via a polyethylene glycol (PEG) linker of appropriate length to provide flexibility and increase the efficiency of the crosslinking reaction between the protein partners. Additionally, a crosslinking substrate may append a fluorophore or affinity ligand of choice. The interacting proteins are genetically fused to self-labeling tags, such as SNAP-tag or CLIP-tag, and specifically crosslinked using a suitable bifunctional substrate [26,29,41]. After in vivo crosslinking, the trapped protein complexes are analyzed by Western blot or in-gel fluorescence scanning. Kai Johnsson and co-workers demonstrated that this approach can detect weak or transient interactions, which are difficult to distinguish by conventional chemical methods, and that the crosslinking efficiency can be used as an indicator of the interaction strength between two proteins [41]. In a complementary approach, Wymann and co-workers developed a crosslinking substrate (HaXS), on the basis of SNAP-tag and HaloTag self-labeling proteins, that functions as a chemical inducer of dimerization and as such can force protein– protein interactions and promote protein translocations to various cellular compartments. This system was shown to trigger PI3K/mTOR signaling pathways, without interfering with endogenous signaling molecules or induction of feedback mechanisms. HaXS dimerizers were also used in conjunction with the rapamycin-based dimerization system for controlled assembly of a multimeric protein complex. Current Opinion in Chemical Biology 2014, 20:36–45

The second type of multifunctional reporter probe incorporates multiple labels in the same substrate molecule (Figure 2d). These are designed for tandem imaging and immobilization/affinity purifications. Lambert and co-workers utilized a SNAP-tag bifunctional substrate that incorporates both a near infrared emitting fluorophore (Dy649) and a biotin-terminated PEG chain (BG-649-PEG-Biotin) to probe self-association of GPCRs in cell membranes [42]. GPCRs play an important role in signal transduction and are targets for a large number of therapeutic drugs. GPCR oligomerization was assessed by recruiting subsets of families A or C protomers into microscopic domains created by micron-sized streptavidin beads on the surface of living cells. This assay is conceptually similar to in vitro co-immunoprecipitation assays, but does not require physical removal of receptors from the plasma membrane. The dimerization of family C metabotropic glutamate receptor 2 (mGlu2) was confirmed by showing that mGlu2 protomers labeled with a fluorophore only (SNAP-green) were corecruited into streptavidin bead domains at the same time as mGlu2 protomers labeled with BG-649-PEG-Biotin. In contrast, only b2 adrenergic receptor (b2AR) protomers labeled with BG-649-PEG-Biotin were recruited to streptavidin bead domains, suggesting that interactions between these family A protomers are too weak to directly influence subcellular location. Another example of an application utilizing multifunctional reporters is for single-molecule imaging of binding and nucleation events of actin filaments in vitro. Gelles and co-workers tethered the actin cytoskeleton protein complex Arp2/3 to a slide surface via the bifunctional BG-649-PEG-Biotin and monitored binding of fluorescently labeled nucleation promoting domains (diVCA) and actin filaments from solution [43].

Recent applications of tag-mediated labeling approaches Superresolution imaging

The last decade has seen tremendous advances in imaging techniques virtually bridging the gap between conventional fluorescence microscopy and electron microscopy [34]. Imaging beyond the diffraction limit of light is now made possible by a vast repertoire of superresolution techniques, including single-molecule localization microscopy (SMLM), stimulated emission depletion (STED) microscopy, and structured illumination microscopy (SIM). SMLM approaches, such as PALM and stochastic optical reconstruction microscopy (STORM), employ multiple cycles of stochastic activation and detection of individual fluorescent molecules in the sample followed by single-molecule localization and image reconstruction. In these methods, the spatial location of single molecules are determined either by photoactivation or photoswitching of fluorescent reporters. Depletion-based approaches, such as STED, overcome the diffraction limit with the aid of two laser beams (an excitation and a depletion laser) that restrict the www.sciencedirect.com

Fluorescent imaging Correˆa 41

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(a) Comparison of confocal versus STED superresolution imaging. Human embryonic kidney (HEK 293) cells stably expressing epidermal growth factor receptor (EGFR)-SNAP were labeled with BG-Chromeo494, washed, and then incubated with the pre-labeled epidermal growth factor ligand EGFCLIP-ATTO647N. Two-color STED imaging of the EGFR-EGR complex resulted in an improvement of 3-fold to 4-fold in resolution. (b) A SNAP-tag based pulse-chase imaging experiment. HeLa cells transiently transfected with centromeric histones N-terminally tagged with SNAP and a human influenza hemagglutinin (HA) epitope tag. Cells are pulse-labeled with a fluorescent substrate (SNAP-Cell TMR-Star). Following substrate washout (chase), the turnover of labeled proteins incorporated into centromeric chromatin can be visualized and quantified at various time points.

spatial area of emission of fluorophores reporters in a selective manner. SIM approaches rely on illumination using grid patterns to extract information from the image focal plane and achieve subdiffraction resolution. The success of any of these techniques for visualization of cellular ultrastructure depends heavily on the optical properties of fluorophores (e.g., strong brightness and resistance to photobleaching) and on the ability to target these fluorophores to specific cellular targets. Small molecule fluorescent reporters have been successfully used in combination with tagging techniques to image cytoskeletal and cell membrane proteins [44], clathrincoated pits [38], mitochondria [35], and histone proteins [45] with nanoscale resolution (Figure 3a). Superresolution imaging of an amebic endoplasmic reticulum [46] and of clusters of the human immunodeficiency virus (HIV-1) Gag protein [47] have also been demonstrated. Several fluorescent reporters have been custom designed to meet the biophysical requirements of superresolution techniques, including silicon–rhodamines (STORM/STED) [33], www.sciencedirect.com

photoactivatable caged rhodamines (PALM) [35], and photoswitchable cyanine dyes (STORM) [48]. Multicolor superresolution imaging has been realized both in live and fixed cells. For instance, two-color STED was utilized to image epidermal growth factor (EGF) and EGFR interaction [49] and to characterize the association of centrosomal proteins [29]. Very recently, Xie and co-workers developed an optical imaging technique that combines reflected lightsheet illumination with superresolution microscopy (RLSSRM), allowing them to probe the spatial organization of RNA polymerase II (RNAP II)-mediated transcription with subdiffraction resolution. Additionally, Xie and co-workers quantified RNAP II clustering in the nucleus with singlecopy accuracy by means of a two-color colocalization experiment, where SNAP-tagged RNAP II molecules were simultaneously labeled with either SiR or TMR dyes [50]. The exceptional spatiotemporal resolution achievable by a combination of site-specific labeling technologies and live-cell superresolution microscopy makes this a powerful approach for structural and mechanistic investigation of biological processes at nanoscale resolution. Current Opinion in Chemical Biology 2014, 20:36–45

42 Molecular imaging

Single-molecule imaging Imaging strategies on the basis of single-molecule fluorescence of labeled target proteins have been designed to track protein dynamics and the assembly of protein complexes in vitro and in living cells. For example, Manley and co-workers have demonstrated multicolor single-molecule tracking of stochastically active fluorophores on the surface and in the nucleus of live cells [51]. In addition to SMLM-based approaches, total internal reflection fluorescence (TIRF) microscopy has been utilized to image proteins at the single-molecule level. TIRF provides excellent temporal resolution for dynamic studies of single molecules due to the exceptionally high signal-to-noise ratio generated by evanescent wave excitation. Lohse and coworkers employed receptor labeling with SNAP-tag and TIRF to dynamically monitor single receptors on the surface of live cells and compare the spatial arrangement, mobility, and supramolecular organization of different GPCR family members, including the b1-adrenergic receptor (b1AR), the b2AR, and the gaminobutyric acid (GABAB) receptor [52]. Jun and coworkers investigated single-molecule diffusion dynamics of Notch receptors on the surface of live cells [53]. Notch plays a central role in cell-fate decisions during development, normal tissue maintenance and cancer, but little is known about the dynamics of this receptor in intact cells. The use of small, modular monovalent quantum dots (mQDs) to label a SNAP-tagged Notch fusion enabled the tracking of individual Notch receptors without perturbing their diffusion along the membrane. Using TIRF-based colocalization single-molecule spectroscopy (CoSMoS), Moore and co-workers observed real-time spliceosome assembly in Saccharomyces cerevisiae whole cell extracts with aid of surface-tethered pre-mRNA molecules and individual spliceosomal subcomplexes fluorescently tagged with TMP-tag and SNAP-tag [54,55]. The combination of CoSMoS with tagging tools was also reported by Gelles and co-workers to examine the dynamics of association and branching of actin networks via three-color single-molecule imaging experiments [43].

Homogenous time-resolved fluorescence Protein–protein interactions are at the core of a vast number of cellular processes. Although there are many methods available to analyze protein–protein interactions, the accurate characterization of interacting protein partners continues to be one of the main challenges in functional proteomics. FRET is one of the most commonly used techniques for studying protein–protein interactions in living cells. However, traditional FRET approaches that utilize fluorescent proteins suffer from their relatively low brightness and broad absorption/emission spectra as well as from the background fluorescence of the cell and other sample components (buffers, chemical compounds, etc.). Site-specific labeling technologies offer the possibility to employ long lifetime emission Current Opinion in Chemical Biology 2014, 20:36–45

lanthanoid cryptates as fluorescence donors. Long-lived fluorophores allow for Homogenous Time Resolved Fluorescence (HTRF) where the short-lived background fluorescence is eliminated by introducing a time delay (approximately 50–150 ms) between the excitation and fluorescence measurement. This approach has been utilized to study the interactions of several membrane protein receptors, including g-aminobutyric (GABAB) [52,56], glutamate [57], dopamine [58], ghrelin and somatostatin receptors [59].

Pulse-chase labeling Direct observation of protein dynamics in living cells is key to understanding their underlying molecular and cellular functions. Self-labeling systems present a unique platform to differentially label protein cohorts with spatial and temporal control. Since the timing of labeling of a given fusion protein is under experimental control, protein trafficking and turnover, macromolecular assembly, and organelle dynamics become open to investigation by means of pulse-chase and quench-pulse-chase labeling experiments [25]. In pulse-chase labeling, the fusion protein is labeled with a fluorescent reporter (pulse) followed by removal of excess probe (optionally a nonfluorescent substrate can be added to terminate and block further labeling). After a given period of time (chase), the fate of the labeled protein at a particular subcellular location is determined without competing signals from newly synthesized fusion proteins (Figure 3b). The SNAP-tag pulse-chase approach was exploited, for instance, to investigate the regulation of a Mycobacterium tuberculosis intramembrane metalloprotease crucial for virulence [60] and to determine the decay rate of cyclin B in Giardia intestinalis [61]. This strategy is easily adaptable and can be used to study protein dynamics in a range of organisms at any time frame (hours, days) post-labeling. For the analysis of newly synthesized proteins only, a block, or quench step is first applied. In this protocol, termed quench-pulse-chase (or quench-chase-pulse), a non-fluorescent substrate is added to the media to label any fusion protein that is present at the onset of the experiment, followed by a given amount of time for nascent protein expression (chase) and labeling with a fluorescent probe (pulse). Only the pool of proteins synthesized during the chase period is fluorescently labeled and thus will be used for imaging and quantitation. For example, Jansen and co-workers employed quench-pulse-chase labeling to probe the roles of centromeric protein A (CENP-A), a histone H3 variant, in centromere determination and cell division [62,63]. In addition, quench-pulse-chase experiments can be combined with cell synchronization to determine the fate of a newly synthesized pool of protein during the cell cycle, and with siRNA-methods to deplete endogenous (untagged) protein expression [64]. Lastly, multicolor pulse-chase labeling experiments can be used to facilitate the differentiation of older and newer copies of a given www.sciencedirect.com

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protein. Solimena and co-workers utilized multicolor pulse-chase labeling to image insulin secretory granule aging in rat insulinoma INS-1 cells, thereby overcoming limitations encountered with strategies on the basis of radiolabeling or fluorescence timer proteins [65].

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Concluding remarks The development of new fluorescent probes and imaging techniques have dramatically accelerated the pace of discovery and will continue to be of fundamental importance to new insights in cellular processes. Research areas such as superresolution and single-molecule microscopy have seen incredible advances and it is not unreasonable to expect that the use of these techniques for live-cell imaging should become routine in the very near future. The application of fluorescent reporters in medical imaging is another area of increasingly growing interest. As a step in this direction, a new generation of highly sensitive nearinfrared and photoactivatable imaging probes has been devised to characterize and monitor biological processes in complex multicellular organisms. The ability to specifically attach fluorescent reporters to individual proteins is key to deciphering the molecular basis of cellular pathways, such as signal transduction and gene regulation. Collectively, site-specific protein tagging strategies have demonstrated great potential for studying and manipulating proteins in living systems and will certainly contribute to the realization of these exciting challenges.

Conflict of interest The author is an employee at New England Biolabs, Inc. SNAP-tag1 is a registered trademark of New England Biolabs, Inc.

Acknowledgements Ariele Hanek and John Buswell for valuable suggestions and critical reading of the manuscript. Don Comb and Jim Ellard for their continued support of basic research at New England Biolabs, Inc.

References and recommended reading Papers of particular interest, published within the period of review, have been highlighted as:  of special interest  of outstanding interest 1.

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