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Proc Natl Acad Sci U S A. Jun 3, 2008; 105(22): 7744–7749.
Published online May 29, 2008. doi:  10.1073/pnas.0803060105
PMCID: PMC2402386
From the Cover
Cell Biology

A drug-controllable tag for visualizing newly synthesized proteins in cells and whole animals


Research on basic cellular processes involving local production or delivery of proteins, such as activity-dependent synaptic modification in neurons, would benefit greatly from a robust, nontoxic method to visualize selectively newly synthesized copies of proteins of interest within cells, tissues, or animals. We report a technique for covalent labeling of newly synthesized proteins of interest based on drug-dependent preservation of epitope tags. Epitope tags are removed from proteins of interest immediately after translation by the activity of a sequence-specific protease until the time a protease inhibitor is added, after which newly synthesized protein copies retain their tags. This method, which we call TimeSTAMP for time-specific tagging for the age measurement of proteins, allows sensitive and nonperturbative visualization and quantification of newly synthesized proteins of interest with exceptionally tight temporal control. We demonstrate applications of TimeSTAMP in retrospectively identifying growing synapses in cultured neurons and in visualizing the distribution of recently synthesized proteins in intact fly brains.

Keywords: protein synthesis, protein turnover, synaptic plasticity, synaptogenesis

Spatially controlled protein production and delivery are fundamental processes in the development, maintenance, and adaptation of specialized cellular structures. Local synthesis allows for the rapid production of proteins in regions of the cell where they are needed. For example, local protein synthesis is associated with myofibril growth in cardiac myocytes (1) and contributes to actin production at the leading edge of migrating fibroblasts (2, 3). During neuronal development, guidance of axons to their targets involves the localized induction of translation in the axonal growth cone by extracellular factors (4, 5). In mature neurons, distal dendrites locally synthesize proteins in response to local stimulation by growth factors or neurotransmitters (68). The induction of long-term potentiation (LTP), an electrophysiological model of learning, induces the redistribution of polyribosomes to synapses and the enlargement of polyribosome-associated synapses (9, 10). Indeed, local dendritic translation is required for establishment of LTP (11). The fragile X mental retardation protein is required for stimulus-induced translation of a subset of dendritic messages, including synaptic structural elements (12, 13), implying that abnormalities in activity-dependent local synthesis of synaptic proteins may underlie some disorders of mental cognition as well.

Delivery of newly synthesized proteins to subcellular regions is also essential in maintaining specialized cellular functions. For example, sorting within the secretory pathway allows for long-distance transport of proteins from the endoplasmic reticulum to discrete final destinations within the cell. This process is necessary for the establishment and maintenance of polarized epithelial cells and of axonal and dendritic specializations in neurons, including presynaptic and postsynaptic complexes (14, 15). Local demands for particular proteins could also contribute to spatially specific protein accumulation. For example, in neurons, synapses undergoing growth could be expected to accumulate recently synthesized structural proteins from nearby pools at higher rates than stable synapses.

Studies of new protein synthesis and delivery, especially in neuronal plasticity, would benefit from the ability to visualize new proteins. However, existing methods are limited either in fidelity for reporting new proteins or compatibility with tissues or living animals. Destabilized fluorescent proteins (FPs) are used as proxy reporters of translational regulation by untranslated regions (UTRs) of mRNAs (16), but destabilized FPs are unsuitable as fusion tags of new protein localization because fusing one to a stable protein of interest would create a molecule that is more stable than the destabilized FP alone and/or less stable than the native protein of interest. In the latter case, destabilization could prevent the protein from reaching its final destination or could disrupt the function of complexes that contain the protein. Turnover of destabilized FPs is also not controllable and depends on proteasome activity, which may vary across conditions. The FP tetramer fluorescent timer slowly shifts emission over time due to fluorescence resonance energy transfer from green to red subunits as red chromophores mature (17), but it has not been used as a protein tag, likely because of its multimeric nature and uncontrollable time course. FPs that can be converted from green to red by intense short-wavelength light, such as Kaede, Eos, and Dendra, allow visual differentiation of proteins synthesized before and after photoconversion (18, 19). Similarly, fluorescence recovery after photobleaching (FRAP) allows selective visualization of newly synthesized proteins. However, photoconversion or FRAP requires uniform delivery of high-intensity UV or violet light throughout the volume of interest, which will be difficult in intact organisms and, in our experience, already causes substantial phototoxicity when applied over entire neurons. FP readouts also do not allow signal amplification and are delayed by chromophore maturation.

We have studied protein trafficking and turnover in cultured cells by sequential application of green and red biarsenical dyes (FlAsH and ReAsH) that bind to tetracysteine tags (20). However, this and similar pulse–chase chemical labeling technologies such as SNAP-tag (21) require stoichiometric binding of the first label followed by rapid washout and introduction of the second, which will be difficult at best in thick tissues and impossible in animals. Incorporation of radioactive or unnatural amino acids labels new proteins (22, 23), but nonspecifically labels all new proteins including abundant high-turnover ones and therefore cannot be used for high-resolution imaging of newly synthesized proteins of interest.

Improvements in technology are therefore necessary before visualization studies of new protein accumulation in local adaptive processes such as synaptic plasticity can be performed in the setting of an entire tissue or organism. The ideal system for visualizing newly synthesized proteins should be highly sensitive and specific for new proteins, controllable in terms of the time periods that can be investigated, applicable in whole animals, and compatible with biochemical and optical detection methods at various spatial scales. We describe a system for epitope tagging of newly synthesized proteins under the control of a cell-permeable drug that can be used in animals. In this method, an epitope tag is rapidly removed from proteins of interest by a sequence-specific protease unless a protease inhibitor is present, whereupon newly synthesized protein copies retain the tag. We call this method TimeSTAMP, for time-specific tag for the age measurement of proteins. We demonstrate the ability of TimeSTAMP (TS) to identify retrospectively new synapses in neurons based on the content of new synaptic proteins and to generate three-dimensional maps of new protein distributions in whole fly brains.


Design of a Drug-Regulatable Epitope Tag.

We desired to develop a tag whose presence on new proteins could be induced by the one-time administration of a small-molecule drug to either cells or animals. The strategy we devised was to incorporate a specific protease activity to confer self-removing behavior onto an epitope tag and then to use a corresponding protease inhibitor to block tag removal, so that only proteins synthesized after inhibitor application would be tagged (Fig. 1A). We identified the hepatitis C virus (HCV) NS3 protease as uniquely suitable for several reasons: the NS3 protease domain is small (19 kDa) and monomeric, demonstrates an unusual but well characterized substrate specificity, can be expressed in mammalian cells without noticeable toxicity, and can be specifically blocked by cell-permeant drugs developed by the pharmaceutical industry such as BILN-2061 (2426).

Fig. 1.
TS concept and proof of concept. (A) Strategy for drug-dependent epitope tagging of newly synthesized proteins. (B) PSD-95 fused to an NS4A/B protease site, an NS3 protease domain with a T54A mutation, another NS4A/B site, and cyan fluorescent protein ...

We performed initial tests of our strategy in cells transfected with the model protein PSD-95 fused to combinations of various versions of NS3 protease, cognate protease sites, and tags [supporting information (SI) Fig. S1A]. NS3 protease lacking the NS4A cofactor efficiently and specifically cleaved at NS4A/B target sites in a manner inhibitable by BILN-2061 (Fig. S1B), with the best inhibition observed on a slow-cleaving NS3 variant with a Ala-54 to Thr mutation (Fig. 1B and Fig. S1C). We therefore designated a cassette comprising the NS4A/B cleavage site from the HCV polyprotein, the NS3 domain, and another NS4A/B site as the TS module, which can be fused in between a protein of interest and an epitope tag. We designate a TS module with the slow-cleaving Ala-54 NS3 variant as TSa and with the fast-cleaving Thr-54 variant as TSt. Due to its symmetric design, the TS module is suitable for tagging at either end of a protein (Fig. S1D). No off-target cleavage was observed in various model proteins by using either the TSa (Fig. 1 B and C and Fig. S1D) or TSt (Fig. S1B, set 4, and S1D) cassette.

We assessed how rapidly tag accumulation occurs after drug application. In cells expressing the activity-regulated protein Arc fused to a TS module and a distal HA tag, HA was completely removed in the absence of drug but was clearly detectable from the first time point (3 min) after drug application (Fig. 1C). A T7 epitope tag placed proximal to the cleavage sites showed that the protein was expressed at all times (Fig. 1C). Initial accumulation of HA-tagged protein followed a linear relationship (R = 0.997), as expected given a constant rate of production (Fig. 1D). The ability to detect tag accumulation within 3 min of drug application demonstrates that the TS strategy allows control of tag preservation with high temporal resolution.

We tested the ability of TS to tag new proteins in primary hippocampal neurons. Cultured neurons expressing PSD-95-GFP fused to TSa and a C-terminal HA tag (PSD-95-GFP-TSa-HA) showed punctate HA immunofluorescence (IF) in the presence of BILN-2061 but not in its absence (Fig. 2A). No HA staining was observed in transfected neurons in the absence of drug, demonstrating that cleaved tag degrades rapidly and/or is lost during fixation (Fig. 2A). Similar results were obtained with TSa fusions to synaptic proteins Arc and Neuroligin1 (Fig. S2A). To recapitulate endogenous mechanisms of mRNA regulation, the 5′ and 3′ UTRs were included in all constructs. As expected, the protein synthesis inhibitor cycloheximide prevented HA signal accumulation (Fig. 2A), whereas brain-derived neurotrophic factor, which induces Arc translation in neurons (27), increased accumulation of Arc-TSa-HA (Fig. S2B). Drug induced a 20-fold increase in signal with TSa and a 121-fold increase with TSt (Fig. 2B). HA staining detected newly synthesized PSD-95-GFP-TSa-HA at levels comparable with endogenous PSD-95 (Fig. 2C). Thus, TS is specific, sensitive, and generalizable.

Fig. 2.
TS allows selective and sensitive labeling of new PSD-95 in neurons. (A) TS-mediated IF is specific for new proteins. Eighteen days in vitro (DIV) neurons at 9 days posttransfection (DPT) with PSD-95-GFP-TSa-HA show synaptic HA IF after 6 h of BILN-2061. ...

We asked whether TS tagging might perturb protein behavior or be toxic. PSD-95-GFP-TSa-HA was present in dendrites and enriched in puncta in dendritic spines, indicating proper localization (Fig. 2A). TS tagging did not significantly affect protein turnover because replacement rates of PSD-95-GFP-TSa-HA in stably transfected neurons were consistent with the previously measured 36-h half-life of endogenous PSD-95 (Fig. 2D) (28). We observed no abnormalities in neuronal morphology or synaptic density after expression throughout synaptogenesis of PSD-95 fused to TSa or TSt modules (Fig. S2C, constructs 4, 6, and 7). Test constructs containing the more active wild-type NS3 domain fused permanently to PSD-95 resulted in lower synaptic densities (Fig. S2C, constructs 1 and 3), but this effect could be rescued by either allowing protease self-release from PSD-95 (Fig. S2C, construct 2), as occurs in the TS modules, or introduction of the T54A mutation (Fig. S2C, construct 5). These experiments demonstrate that the use of TS modules to label proteins is nonpertubative and nontoxic.

We investigated whether TS can report the spatial distribution of new proteins. In neurons expressing PSD-95-GFP-TSa-HA treated with BILN-2061 for 6 h, a time in which ≈11% of PSD-95 molecules will turn over, assuming steady-state expression and a 36-h half-life, we observed gradients of new protein down the dendrites, distinct from total PSD-95-GFP, which was distributed throughout the cell (Fig. 2E). Likewise, we observed new Neuroligin1 predominantly in the soma after 6 h, distinct from total Neuroligin1 (Fig. S2D), consistent with processing in the secretory pathway. To verify these findings by using another method, we expressed PSD-95 or Neuroligin1 fused to tandem dimer EosFP, which converts from green to red emission upon near-UV irradiation (19). After photoconversion of existing protein into red fluorescence, new protein appears green. Although photoconversion causes death of some neurons, it allowed us to confirm in surviving neurons that newly synthesized PSD-95 and Neuroligin1 exist in a gradient from the soma (Fig. S3). These results show that the TS method can report distributions of newly synthesized proteins.

Application of TS to Tracking Synaptic Growth.

Growth or de novo formation of synapses occurs in response to circuit activity or biochemical stimulation and may underlie some forms of learning (9, 2931). Determining where synapses are growing is therefore critical in understanding circuitry adaptation. Although time-lapse microscopy can track synaptic growth in sparsely labeled neurons in superficial brain regions (32), visualizing growth in deep regions or during unrestrained behavior has not been possible. A plausible strategy may be to image new protein accumulation by growing synapses. A large body of evidence suggests that PSD-95 accumulates in growing synapses from a diffuse cytoplasmic pool (33). Furthermore, based on local photoactivation and FRAP experiments, spines in differentiating hippocampal neurons appear to contain two populations of PSD-95 molecules, with 25–40% of PSD-95 protein exchanging with a half-time of minutes and the remainder relatively immobile on the order of hours (3436).

We explored the possibility of using TS on synaptic proteins to assess synaptic growth retrospectively over a few hours. We acquired GFP images of neurons expressing PSD-95-GFP-TSa-HA during synaptic differentiation in the presence of BILN-2061 for 6 h, a time interval appropriate for detecting the formation of several synapses per microscope field (37). Comparing GFP images from the beginning and the end, we identified new postsynaptic densities (PSDs) as newly appearing PSD-95-GFP puncta that colocalized with the presynaptic marker synapsin. These new PSDs showed significantly higher HA IF relative to GFP, indicating that they preferentially incorporate new PSD-95 (Fig. 3A). New PSDs were also smaller on average (Fig. 3A), but small size alone is not a sufficient indicator of new synapses because some neurons exhibited small PSDs at the end of the experiment that existed before the observation period (Fig. 3B). These stable small PSDs also had HA/GFP ratios similar to those of nearby stable large PSDs (Fig. 3B), demonstrating that larger PSDs do not inherently demonstrate lower HA/GFP ratios, e.g., because of incomplete antibody access. These results show that a high fractional content of new PSD-95 characterizes recently formed synapses. Using 3-h intervals, we confirmed that the HA/GFP ratio was correlated with PSD newness, so that more recently appearing PSD had significantly higher HA/GFP ratios than those appearing earlier (Fig. 3C), further supporting this relationship and demonstrating that differences in new PSD-95 content arise within 3 h. In all of our experiments, we observed that old PSDs maintained distinctly lower HA/GFP ratios than the adjacent dendritic shaft, confirming that synaptic PSD-95 molecules are not in complete rapid exchange with the PSD-95 pool in the dendritic shaft. Taken together, these results show that visualization of newly synthesized proteins by TS can be used to identify nascent synapses retrospectively in differentiating neurons.

Fig. 3.
Newly forming synapses preferentially accumulate new PSD-95. (A) Twenty-one days in vitro neurons at 14 DPT expressing PSD-95-GFP-TSa-HA were imaged at the time of BILN-2061 addition. After fixation 6 h later, cells were stained for HA for newly synthesized ...

Whole-Brain Mapping of New Protein Distributions in Living Animals.

The ability of TS to be controlled by a cell-permeable drug should allow time-specific protein tagging in a living animal. The fruit fly Drosophila melanogaster has emerged as a preeminent model for studying neuronal circuitry (38). We chose to examine fly calcium/calmodulin-dependent protein kinase II (dCaMKII), whose transcription occurs throughout the nervous system (39) but whose translation and synaptic localization are influenced by neuronal activity (40). We asked whether TS could be used to generate a three-dimensional map of new dCaMKII protein to reveal whether new dCaMKII distributions vary between or within neurons of the fly brain. Because the dCaMKII C terminus mediates homododecamerization, we tagged dCaMKII at the N terminus by using a TSt module in which an HSV tag serves as the a drug-dependent epitope and a HA tag as a constitutive epitope. This HSV-TSt-HA-dCaMKII fusion protein demonstrates BILN-2061-dependent HSV tag preservation in transfected cells at the ambient temperatures in which flies are raised (Fig. S1D). We then created transgenic fly lines expressing HSV-TSt-HA-dCaMKII under the control of the elav promoter, which drives expression in all neurons with enrichment in the mushroom bodies (MBs), a pattern similar to endogenous dCaMKII protein (39). We administered BILN-2061 for 6 h to living flies, then we stained and visualized HSV-tagged new and HA-tagged total protein by confocal microscopy. HA revealed that total HSV-TSt-HA-dCaMKII protein was expressed throughout brains with enrichment in the MBs, similar to elav-driven tubulin-GFP (Fig. 4A). As expected, HSV IF was observed only in the presence of inhibitor (Fig. 4A).

Fig. 4.
TS tagging of newly synthesized proteins in whole flies. (A) Adult flies expressing HSV-TSt-HA-dCaMKII and tubulin-GFP in neurons were administered BILN-2061 for 6 h, then brains were stained. HSV IF reveals distributions of newly synthesized dCaMKII, ...

New dCaMKII was particularly abundant in scattered neurons in the anterior protocerebrum near the MB lobes and in groups of Kenyon cells (KCs) of the MB (Fig. 4A and Fig. S4A). KCs project axons through the peduncles that then branch in the α, α′, β, β′, and γ lobes of the MBs. High levels of new dCaMKII could be traced from the KC soma along discrete axonal bundles continuous with moderate levels of new dCaMKII in the α and β lobes (Fig. 4 B and C and Fig. S4A). In contrast, no HSV IF was observed in the α′, β′, and γ lobes, even though total dCaMKII protein, as revealed by HA IF, is equally abundant in the α, α′, β, β′, and γ lobes (Fig. 4C). These results indicate that KCs projecting to the α and β lobes maintain higher rates of dCaMKII synthesis than KCs projecting to the α′, β′, and γ lobes. Interestingly, α/β KCs represent a developmentally and functionally distinct population of KCs (41, 42).

TS tagging also revealed subcellular differences in new protein distributions. Within KCs, HSV intensities and HSV/HA ratios are complementary to total dCaMKII, with higher levels in the cell bodies and peduncle than in the distal axon branches (Fig. 4). This pattern of dCaMKII turnover is consistent with dCaMKII production in the KCs occurring predominantly in soma. Similar results were obtained in flies expressing the slower cleaving HSV-TSa-HA-dCaMKII (Fig. S4B). Taken together, these results show that the TS technique reveals heterogeneity in dCaMKII production between neurons and in the subcellular distribution of new dCaMKII molecules in the brains of living animals.


By tightly linking onset of protein tagging to the presence of a drug, the TS method allows temporally controlled labeling of newly synthesized proteins of interest in thick tissues or intact animals. We have demonstrated that TS is sensitive, specific, nonperturbing, nontoxic, and generalizable. Taking advantage of these attributes, we have demonstrated that growing synapses in primary neurons preferentially accumulate new PSD-95 molecules. We have also visualized the distribution of newly synthesized dCaMKII throughout the fly brain, demonstrating that TS can be used to label proteins synthesized during unrestrained behavior in animals.

In addition to its unique ability to control protein tagging in vivo, TS combines the temporal resolution of small-molecule regulation and the high sensitivity and specificity of antibodies, and it may be the method of choice in some in cellulo situations as well. Signal can be detected within minutes of drug addition, a time resolution finer than sequential chemical labeling with its long incubation and wash times. The sensitivity of antibody staining, where signal amplification can be achieved with secondary antibodies, allows the visualization of even low-abundance proteins. For example, synaptic proteins such as PSD-95 are present at a few hundred copies per synapse (43, 44), only a fraction of which will be synthesized within a few hours. In contrast, signals from photoconvertible proteins cannot be further amplified, are limited by photobleaching of converted protein, and are delayed by chromophore maturation.

TS should be adaptable to other antibody-based detection methods. For example, immunoelectron microscopy could be performed to reveal new proteins at the synapse with ultrastructural resolution. The TS method also allows multiplexing; unique epitope tags on different proteins of interest can be detected with antibodies conjugated to different fluorophores for fluorescence microscopy or to differently sized quantum dots for correlated fluorescence and electron microscopy (45). We have already demonstrated TS in immunoblotting, where it allows quantitation of proteins produced before vs. after a time of interest. It has also not escaped our attention that highly specific protease–inhibitor pairs could be used to regulate the attachment of functional peptide motifs, not just epitope tags, and work is needed to explore these possibilities.

Our results with PSD-95 also provide proof of concept for an approach to identifying synapses undergoing growth within the intact brain in response to environmental changes, learning, or pathway stimulation. Although we have focused on postsynaptic proteins, presynaptic proteins will also be of interest. Other proteins may be more immobile or turn over more slowly in stable synapses than PSD-95. As candidates are identified, it will be of interest to test their ability to serve as reporters of synaptic growth.

Materials and Methods

Construction and Initial Testing of TS.

HCV genotype 1a NS4A and NS3 coding sequences were obtained by RT-PCR from infected macaque liver RNA. The HCV NS3 protease inhibitor BILN-2061 was custom-synthesized by Acme Biosciences. For initial tests by immunoblotting, linear fusions of PSD-95, the HCV NS3 protease domain lacking the helicase domain flanked on both sides by cleavage sites, an HA epitope tag, and the CFP were constructed. The NS4A β strand, which enhances NS3 protease activity in trans (46), was either fused N-terminal to the NS3 domain, a configuration shown to possess high catalytic activity (47), or omitted. In some constructs and in the final TSa cassette, we introduced a mutation of Thr-54 to Ala that had been found to reduce the catalytic rate of the enzyme 10-fold (48). Analysis of the protease structure (49) revealed Thr-54 to be distant from the catalytic triad and the BILN-2061 binding site. Rather, Thr-54 may help shape the oxyanion hole; its hydrogen bond with the backbone oxygen of Leu-44 may help orient the Leu-44 side chain, which in turn interacts with the peptide backbone of the oxyanion hole. For cleavage sites, we chose the NS4A/4B junction because it is efficiently cleaved by NS3 protease and because the NS4A cleavage product can bind to the active site and competitively inhibit the protease with an inhibitory constant of 0.6 μM (50), possibly limiting further protease activity. Fusions using cleavage sites from the NS5A/5B junction showed reduced sensitivity to inhibition by BILN-2061 (data not shown), consistent with the faster activity of NS3 protease on this sequence (51). We also created for comparison constructs without a cleavage site in between PSD-95 and the protease domain. HEK293 cells were transfected by using Lipofectamine 2000, then grown for 2 days with or without BILN-2061. SDS lysates were prepared, followed by SDS/PAGE and immunoblotting with anti-PSD-95 to differentiate uncleaved and cleaved products by size. Constructs containing the NS4A β strand were inefficiently inhibited by BILN-2061 in early experiments and were not explored further.

Neuronal and Fly Materials and Methods.

See SI Materials and Methods.

Supplementary Material

Supporting Information:


We thank Michael McKeown, Holli Weld, and Evangeline Mose for assistance with experiments; Paul Steinbach with microscopy, Stefan Wieland and Frank Chisari (Scripps Research Institute, La Jolla, CA) for liver RNA from HCV-infected macaques, Joerg Wiedenmann (University of Ulm) for EosFP cDNA, Cynthia Hughes, Marco Gallo, Jing Wang, and Charles Zuker (University of California, San Diego, CA) for fly stocks and advice, Terunaga Nakagawa for discussion, and members of the Tsien and Glenn laboratories for their support. This work was supported by a Jane Coffin Childs fellowship (to M.Z.L.), by the Howard Hughes Medical Institute and National Institutes of Health (to R.Y.T.), and by a Burroughs Wellcome Fund Clinical Scientist Award in Translational Research and National Institutes of Health (to J.S.G.).


The authors declare no conflict of interest.

This article contains supporting information online at www.pnas.org/cgi/content/full/0803060105/DCSupplemental.


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