Cellular Incorporation of Unnatural Amino Acids and Bioorthogonal

Review pubs.acs.org/CR

Cellular Incorporation of Unnatural Amino Acids and Bioorthogonal Labeling of Proteins Kathrin Lang*,† and Jason W. Chin* Medical Research Council Laboratory of Molecular Biology, Francis Crick Avenue, Cambridge CB2 0QH, United Kingdom 4.3. Cell-Selective Cotranslational Labeling of Proteins 5. Site-Specific Incorporation of Unnatural Amino Acids for Labeling Proteins 5.1. Current Labeling Approaches 5.1.1. Fluorescent Proteins 5.1.2. Self-Labeling Tags 5.1.3. Self-Labeling Enzymes 5.1.4. Enzyme-Mediated Labeling 5.2. Incorporation of Fluorescent Unnatural Amino Acids 5.3. Incorporation of Unnatural Amino Acids with Bioorthogonal Handles 5.3.1. Incorporation of Ketones 5.3.2. Incorporation of Azides 5.3.3. Incorporation of Alkynes 5.3.4. Incorporation of 1,2-Aminothiols 5.3.5. Incorporation of Reaction Partners for Pd-Catalyzed Cross-Coupling 5.3.6. Incorporation of Photoclick Reaction Partners 5.3.7. Incorporation of Anilines 5.3.8. Incorporation of Tetrazines and Dienophiles for Inverse-Electron-Demand Diels−Alder Reactions 6. Conclusions and Future Challenges Author Information Corresponding Authors Present Address Notes Biographies References

CONTENTS 1. Introduction 2. Chemoselective Reactions in Biology 2.1. Ketone/Aldehyde−Hydrazide/Alkoxyamine Reactions 2.2. Azide−Phosphine Reactions (Staudinger Ligations) 2.3. Azide−Alkyne Reactions ([3 + 2] Cycloadditions) 2.4. Cyclooctynes−1,3-Nitrone Dipoles ([3 + 2] Cycloadditions) 2.5. Strained Alkene−Azide Reactions ([3 + 2] Cycloadditions) 2.6. Alkene−Tetrazole Reactions (Photoclick Cycloadditions) 2.7. 1,2-Aminothiol Condensations 2.8. Alkenes in Olefin Metathesis 2.9. Palladium-Catalyzed Cross-Coupling Reactions 2.10. Other Cycloadditions 2.11. Strained Alkene/Alkyne−Tetrazine Reactions (Inverse-Electron-Demand Diels− Alder Cycloadditions) 3. Incorporation of Unnatural Amino Acids into Proteins 3.1. Residue-Specific Incorporation of Unnatural Amino Acids via Selective Pressure Incorporation 3.2. Site-Specific Incorporation of Unnatural Amino Acids via Genetic Code Expansion 4. Applications of Residue-Specific Unnatural Amino Acid Incorporation and Chemoselective Modification 4.1. Global Labeling of Residue-Specifically Incorporated Unnatural Amino Acids 4.2. Profiling Time-Resolved Protein Synthesis (BONCAT and FUNCAT) © 2014 American Chemical Society

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1. INTRODUCTION In the past 15 years, there have been rapid developments in the ability to cotranslationally incorporate unnatural amino acids into proteins produced in cells, both at defined sites via genetic code expansion and throughout the proteome via residuespecific incorporation. There have also been significant and rapid developments in chemoselective reactions for labeling biological molecules in vitro and in cells initiated by the articulation and development of so-called bioorthogonal chemistries. The ability to label proteins with a diverse range of probes has been realized by the cellular cotranslational incorporation of unnatural amino acids bearing functional groups that can be

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Figure 1. Chemoselective labeling of biomolecules. Two-step approach for chemoselective biomolecule labeling: In the first step, a functional group (orange circle), covalently attached to a substrate (light blue rectangle), is introduced into a biomolecule of interest by genetic or chemical methods. In a second step, the functional group reacts chemoselectively, in the presence of all the functional groups found in a living cell or animal, with an added chemical probe (orange arc with red star). Gray shapes denote other biomolecules that must not react with the chemical reporter or the external chemical probe.

labeled using chemoselective reactions. Here, we describe the range of bioorthogonal reactions that have been reported (section 2). We then describe methods for the cellular cotranslational incorporation of unnatural amino acids bearing bioorthogonal functionalities into proteins via residue-specific (section 3.1) or site-specific (section 3.2) approaches. We proceed to outline the application of residue-specific incorporation and labeling (section 4) and the development of sitespecific incorporation and labeling (section 5). Finally, we describe the challenges and opportunities for the cellular cotranslational incorporation of unnatural amino acids and chemoselective labeling (section 6).

Because biological systems operate far from equilibrium and biological processes can be very rapid, the intrinsic kinetics of the reactions used to label biomolecules are very important. Many reactions used to selectively label biomolecules follow second-order kinetics, and their rates depend directly on the concentration of both the biomolecule and the labeling reagent as well as the intrinsic second-order rate constant k2 (M−1 s−1) of the reaction; this makes labeling of low abundance biomolecules, including many proteins at their native abundance, particularly challenging. In principle, low abundance biomolecules may be labeled rapidly by using a large excess of labeling reagent.2 However, this is commonly problematic, because (i) labeling reagents are often not soluble at high concentrations, (ii) labeling reagents are not infinitely specific and participate in off-target reactions with functional groups present at high abundance, and (iii) noncovalent interactions lead to noncovalent labeling of offtarget molecules. The intrinsic second-order rate constant of labeling reactions are therefore a very important variable for biomolecule labeling, and faster chemoselective reactions will enhance the utility of approaches to label proteins. Until recently, however, most of the established chemoselective reactions, for which one partner can be incorporated into a biomolecule, use reagents or catalysts with suboptimal biological compatibility and/or proceed with sluggish rate constants in the range of 10−4−10−1 M−1 s−1 (see detailed discussion in sections 2 and 5.3).1b,3 These rates are sufficient to label purified biomolecules in vitro and to allow useful labeling of metabolically incorporated reporter-bearing glycan analogues presented at high density on the cell surface, as well as labeling of unnatural amino acid analogues incorporated nonspecifically throughout the proteome.4 In general, however, faster and quantitative labeling reactions would facilitate the widest range of applications, and for labeling proteins at defined sites both in vitro as well as in the context of living systems, reactions that are chemoselective, nontoxic, and rapid in a biological context are needed. In recent years, there has been great progress in accelerating chemoselective reactions to facilitate rapid protein labeling in live cells. In sections 2.1−2.11, we review chemistries that have found applications in living systems to label biomolecules.

2. CHEMOSELECTIVE REACTIONS IN BIOLOGY To study the functions and dynamics of biological molecules, methods to covalently and selectively label biomolecules have emerged.1 A two-step approach for biomolecule labeling is commonly employed: (i) a unique chemical functionality is incorporated into a biomolecule of interest either by genetic or chemical methods or by hijacking a metabolic pathway; (ii) an external chemical probe is introduced that reacts with the incorporated chemical functionality in a selective and specific manner (Figure 1).1a,b This approach, in principle, has the great advantage that a single chemical reaction can be used to install a diverse array of probes in a biomolecule. The development and application of suitable chemoselective reactions has however proven to be incredibly challenging. It requires that (i) the incorporated functionality and the probe react selectively with each other under physiological conditions, without either of them cross-reacting with the many other chemical functional groups found in proteins or other biomolecules, (ii) the reactions yield stable covalent linkages with no or innocuous byproducts, and (iii) the reactants are kinetically, thermodynamically, and metabolically stable before the reaction and not toxic to living systems.1a,b Reactions that meet most of these criteria are often described as “bioorthogonal”. Despite the challenges of meeting these criteria, a number of chemoselective reactions have been successfully developed that show good biocompatibility and selectivity in living systems (see sections 2.1−2.11).1a−c However, while a range of reactions are loosely described as bioorthogonal, some of these reactions are chemoselective with respect to many but not all functional groups found in biology. Furthermore, some reactions can only be used to label biomolecules in vitro or on the cell surface, while others have been demonstrated inside cells or intact animals.1a,b

2.1. Ketone/Aldehyde−Hydrazide/Alkoxyamine Reactions

Aldehydes and ketones are two of the most versatile groups in synthetic organic chemistry and were among the first functionalities to be explored as bioorthogonal chemical reporters. While aldehydes react with amines and are used as fixing agents for cells, ketones are less reactive toward the functional groups found in proteins and other biomolecules 4765

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Figure 2. Reaction of ketones/aldehydes with hydrazines or alkoxyamines. (a) Acid-catalyzed condensation of an aldehyde/ketone with reactive amino nucleophiles. (b) Aniline-catalyzed condensation of an aldehyde/ketone with hydrazines or alkoxyamines. (c) Reaction of an aldehyde with a tryptamine nucleophile through a Pictet−Spengler ligation.

sections 4 and 5.3). Furthermore, both engineered ligases (biotin and lipoic acid ligases)15 and aldehyde tags,2,16 as well as chemical methods (periodate oxidation of N-terminal serine or threonine residues17 and pyridoxal phosphate-mediated Nterminal transamination to yield α-ketoamines or glyoxyamides),18 have been developed to modify proteins with ketone- and/or aldehyde-containing handles. In general, ketone/aldehyde reactions with nucleophiles are best suited for in vitro or cell surface labeling,5,12,15a,19 because the reaction commonly requires an acidic pH (4.0 to 6.5), which is difficult to obtain inside most cellular compartments. Furthermore, ketone/aldehyde reporters face competition inside living cells with carbonyl-bearing metabolites like pyruvate, oxaloacetate, sugars, and various cofactors. Moreover, the ketone/aldehyde−hydrazide/hydroxylamine reaction shows quite slow kinetics,20 proceeding with second-order rate constants in the range of 10−4−10−3 M−1 s−1.5,8 To achieve good labeling of the ketone-modified biomolecule, high concentrations (often in the millimolar range) of the labeling reagent are required. This may lead to off-target reactivity and can be problematic in terms of toxicity if applied to labeling within living cells. To overcome the unfavorable acidic conditions and slow kinetics of the ketone/aldehyde−hydrazine/alkoxyamine condensation, aniline was used as a nucleophilic catalyst.21 Aniline considerably accelerates the reaction by forming a highly reactive protonated electrophile with the carbonyl group, which then undergoes rapid transamination to form the hydrazone or oxime product (Figure 2b). Second-order rate constants of 170 and 8.2 M−1 s−1 were reported for the hydrazone and oxime formation, respectively. The reactions were also successfully applied for biomolecule labeling on the cell surface as well as for labeling of a intracellular bacterial receptor.22 Recently, another variant of the aldehyde/hydrazine condensation has been reported.23 To label proteins in vitro, a Pictet−Spengler ligation strategy was used, based on the classic Pictet−Spengler reaction between aldehydes and

under physiological conditions. The synthetic accessibility and small size of ketones have made them amenable to enzymatic incorporation into biological systems through the feeding of close analogues of natural substrates bearing ketones.5 Under acidic conditions (pH 4−6), the carbonyl group of aldehydes and ketones is protonated and reacts with amines to form a reversible Schiff base, the equilibrium usually favoring the free carbonyl form.6 The use of strong α-effect nucleophiles, such as hydrazines or alkoxyamines, however, shifts the equilibrium in favor of the hydrazone and oxime products (Figure 2a).7 The chemoselectivity of the aldehyde−hydrazide/hydroxylamine reaction in living cells was described in 1986 for in situ drug assembly in cancer cells.8 The authors exploited the reactivity between decanal and N-amino-N′-1-octylguanidine, both independently harmless to cells, to form a hydrazonelinked detergent capable of lysing cultured erythrocytes. A similar approach was used to generate a hydrazone-linked inhibitor of proteinase kinase C in situ by assembly of aldehyde and hydrazide based precursors.9 Early examples of using ketones for biomolecule labeling focused on labeling cell surface glycans. An unnatural derivative of N-acetyl-mannosamine bearing a ketone group was synthesized and then metabolically converted to the corresponding sialic acid, which gets incorporated into cell surface exposed oligosaccharides, resulting in the cell surface display of ketone groups. These ketones could then be labeled with hydrazide-based probes under physiological conditions.5,10 Sialic acid analogues bearing ketone groups were also directly fed to cells and incorporated into cellular glycans.11 More recently, ketone reporters were incorporated metabolically into oligosaccharides in bacterial cell walls and labeled with hydrazide-based fluorophores.12 To exploit the ketone−hydrazide/alkoxyamine reaction for chemoselective protein labeling, ketone-bearing groups have been incorporated in the form of unnatural amino acids both in a residue-specific13 as well as a site-specific manner14 (see 4766

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Figure 3. Chemoselective labeling via Staudinger ligation of azides and triarylphosphines. (a) Staudinger reduction between an azide and a triphenylphosphine to yield primary amines. (b) The Staudinger ligation for labeling biomolecules. (c) In the traceless Staudinger ligation, the final amide-linked product is released from the phosphine oxide moiety.

tryptamine nucleophiles.24 The ligation exploits the reaction between an aldehyde tagged protein and alkoxyamines to form an intermediate oxyiminium ion (in an acidic environment), which then undergoes intramolecular C−C bond formation by nucleophilic attack of the indole moiety to form a hydrolytically stable oxacarboline product (Figure 2c). In conclusion, ketones and to some extent also aldehydes have proved to be popular choices as chemical handles for chemoselective biomolecule labeling; however, they should be considered “biorestricted” chemical reporters,1c because they can only be used in certain environments and under specific, often nonphysiological conditions (acidic pH).

Because the azaylide hydrolyzes quickly and breaks the covalent bond between the reactants, the reaction is not a bioconjugation reaction. However, the phosphine reagent has been modified to contain an electrophilic trap (an ester group positioned adjacent to the phosphorus atom); this directs the aza-ylide through a new intramolecular pathway, which competes with destructive hydrolysis.32 The aza-ylide reacts with the electrophilic ester carbonyl group through intramolecular cyclization to form a five-membered ring that undergoes hydrolysis to form a stable amide bond (Figure 3b). This modified Staudinger reaction is now referred to as Staudinger ligation, because it covalently links two molecules together.32a Kinetic analysis of the Staudinger ligation revealed rather slow second-order rate constants in the range of 10−3 M−1 s−1, with the rate-limiting step being the initial nucleophilic attack of the phosphine on the azide nitrogen.31 Attempts to increase the nucleophilicity of the triarylphosphine by adding electron-donating groups to the phosphine failed because these phosphines were more prone to oxidation.1a,33 Shortly after the report on Staudinger ligation, a new variant of this reaction was described, referred to as “traceless Staudinger ligation”, where the final amide-linked product is freed from the phosphine oxide moiety.34 To this end, different phosphine reagents were designed, in which the acyl group is attached via a cleavable linker to the phosphine group, such that once the aza-ylide is formed, attack of the aza-ylide nitrogen on the acyl-carbonyl displaces the cleavable linker and the phosphonium group. Hydrolysis then affords the amide-linked ligation product and liberates the corresponding phosphine oxide (Figure 3c). In a parallel study, a similar approach was

2.2. Azide−Phosphine Reactions (Staudinger Ligations)

In contrast to aldehydes and ketones, the azide group is essentially absent from biological systems and truly orthogonal to the majority of biological functionalities.25 Furthermore, the azide group is small in size and kinetically stable under physiological conditions. These factors make azide-functionalized analogues of metabolic precursors good vehicles for directing the incorporation of azides into numerous biomolecules, including glycans,26 proteins,27 lipids,28 and nucleic acids29 via biosynthetic pathways. In one of the classic reactions employing organic azides, the azide group reacts with phosphines to form primary amines, a reaction known as the Staudinger reduction.30 The reaction proceeds via a nucleophilc attack of the phosphorus atom on the terminal electrophilic nitrogen of the azide to form a phosphazide intermediate that rapidly extrudes N2 to yield an aza-ylide. Under aqueous conditions, the aza-ylide hydrolyzes to yield a primary amine and a phosphine oxide (Figure 3a).31 4767

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Figure 4. Copper-catalyzed azide−alkyne 1,3-dipolar cycloadditions (CuACC) for labeling biomolecules. (a) An azide reacts with a terminal alkyne in a Cu(I)-catalyzed reaction to yield stable 1,4-disubstituted 1,2,3-triazoles. (b) Proposed mechanism for CuACC. (c) Water-soluble ligands BTTES, THPTA, and BTTAA for coordination of Cu(I). (d) Use of copper-chelating organic azides, such as picolyl azides, raises the effective Cu(I) concentration at the reaction site. (e) Strain-promoted azide−alkyne 1,3-dipolar cycloadditions (SPAAC) for biomolecular labeling. (f) Strained cycloalkyne derivatives employed in SPAAC. (g) A light-induced version of SPAAC.

2.3. Azide−Alkyne Reactions ([3 + 2] Cycloadditions)

developed to link a thioester-modified protein to an azidecontaining peptide using phosphinothiol.35 Staudinger ligations have been used to label biomolecules in vitro and in many different cellular environments.36 Cell surface glycan analogues bearing azides were reacted with a watersoluble biotin-containing phosphine reagent in living Jurkat cells, and this chemistry proved to be amenable for use in living animals.32a,37 Azides have also been site-specifically14a or residue-specifically4c,13c incorporated into proteins (see sections 4 and 5) as part of unnatural amino acids, and these azidetagged proteins have been labeled in vitro with phosphinefluorophore or phosphine-polyethyleneglycol (PEG) conjugates via the Staudinger ligation.38 One limitation of the Staudinger ligation is its slow kinetics: the reaction proceeds with a second-order rate constant in the low 10−3 M−1 s−1 range.31 The slow rate of this reaction necessitates the use of high concentrations of the labeling reagent, the phosphine compound, which may generate a high background signal in imaging applications. Attempts to synthesize fluorogenic phosphine reagents based on fluorescein or coumarins addressed the problem to some extent;33 the slow intrinsic kinetics of the reaction, however, remains a limitation. Another drawback of the Staudinger ligation is the oxidation sensitivity of phosphines, which means that phosphine reagents have to be used at relatively high concentrations. Other potential side reactions include the reduction of azides to primary amines by endogenous thiols or other reductants and the potential cross-reactivity of phosphines with disulfides.

Azides can also act as 1,3-dipoles in [3 + 2] cycloadditions with alkynes to yield stable triazoles (Figure 4a). The reaction was first described and examined 50 years ago.39 Although the reaction is highly favorable thermodynamically (ΔG0 ≈ 60 kcal/mol), it requires high temperature and pressure to form the triazole product in reasonable yield.40 In 2002, however, it was reported that the formation of aromatic triazoles starting from terminal alkynes and azides can be catalyzed by copper(I) salts.41 Furthermore, the copper catalyst greatly enhances the regioselectivity of the reaction, yielding predominantly 1,4-disubstitued 1,2,3-triazoles.42 The copper-catalyzed reaction proceeds with rate constants 6−7 orders of magnitude faster than the uncatalyzed reaction. This rate acceleration was mechanistically rationalized by the formation of a proposed copper acetylide to activate terminal alkynes for reaction with azides to form a six-membered Cu(III) metallacycle intermediate (Figure 4b).40,43 The Cu(I)catalyzed reaction, now widely called “the click reaction”,44 proceeds faster in water than in organic solvents and uses functionalities, azides and alkynes, that are generally stable under physiological conditions. Cu(I)-catalyzed click chemistry has been used widely in many different biological studies, for example, to label azides in functionalized viruses,43 to modify proteins in vitro and in vivo,13c,14a,45 and to tag azides in lipids46 and nucleic acids.29c,47 Genetic approaches have been developed to incorporate either the azide group or the alkyne 4768

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successfully for labeling azide-bearing glycans on living cells without any observable cytotoxicity.26b,58 Further synthesis of more reactive cyclooctyne compounds including difluorocyclooctyne (DIFO) derivatives, 59 dibenzocyclooctynes (DIBO), 6 0 and biarylazacyclooctynone compounds (BARAC)61 led to faster reactions with azides (Figure 4f). These probes are more reactive toward azides than the firstgeneration cyclooctyne compounds and have been used to probe azide-containing biomolecules within complex biological systems, including live mammalian cells, C. elegans, and zebrafish embryos.26b,c Despite the rate improvement, the most reactive of the strained cyclooctyne derivatives showed rate constants from ∼0.1 to 1 M−1 s−1. Furthermore, the highly demanding and often low-yielding synthesis of the cyclooctyne probes makes probe development challenging;26b,59 several cyclooctyne probes are now, however, commercially available. More recently, thiacycloheptynes (e.g., 3,3,6,6-tetramethylthiocycloheptyne, TMTH, Figure 4f) have been developed as a new class of reagents for Cu-free SPAAC.62 By masking the triple bond in dibenzocycloctynes as cyclopropenones,63 a light-induced version of SPAAC has been developed. The cyclopropenones do not react with azides in the dark; irradiation with wavelengths around 350 nm, however, generates the corresponding dibenzocyclooctynes via the extrusion of CO, that undergo cycloaddition with azides to give the corresponding triazoles (Figure 4g). Light activation of cyclopropenones has been used to label azido-modified sugars at the cell surface of mammalian cells in a temporally controlled manner,63d and has the potential to facilitate spatially controlled labeling.

group into proteins in the form of unnatural amino acids (see sections 4 and 5).4c,13c,14a The copper-catalyzed reaction of azides with alkynes proceeds with rate constants at least 25 times faster than the Staudinger ligation of azides with triarylphosphines in cell lysates in physiological settings.3,48 However, the main disadvantage of click chemistry is its reliance on the copper(I) catalyst, which may be toxic to cells.49 This apparent toxicity stems from Cu(I)-mediated generation of reactive oxygen species (ROS) from O2.50 For example, E. coli expressing an azide-modified protein are no longer able to divide after 16 h of exposure to 100 μM CuBr, and mammalian cells tolerate low Cu(I) concentrations (600 proteins in primary resting T cells activated by stimuli.149 Newly synthesized proteins that cotranslationally incorporate unnatural amino acids bearing azides or alkynes can also be directly labeled with azide or alkyne functionalized fluorescent dyes. This approach, dubbed FUNCAT (fluorescent noncanonical amino acid tagging), has been demonstrated in both bacterial and mammalian cells (Figure 23) and allows labeled proteins to be visualized by fluorescence microscopy or gelband imaging methods.126b,150 By pulse-labeling mammalian cells with two reactive methionine analogues, Aha and Hpg, and reacting them with two distinct fluorescent dyes, coupled to either azide or alkynyl functionalities, two temporally defined protein populations could be labeled in fixed mammalian cells (Figure 24). The power of BONCAT and FUNCAT approaches lies in the potential to distinguish or separate newly synthesized proteins from the preexisting protein pool. This enrichment permits analysis and identification of protein synthesis in response to internal and external stimuli. Because cells may respond to changing environmental conditions by altering the ensemble of proteins they express, BONCAT can address comparisons of two or more protein ensembles in different biological states, for example, the cancerous versus the noncancerous state. Furthermore, by enriching a subset of the proteome, sample complexity is decreased, which might enable identification of newly synthesized proteins expressed at low

levels, which are difficult to detect by more conventional methods. BONCAT and FUNCAT techniques have been successfully applied to study the proteome of mammalian cells,126a,146 as well as to visualize newly synthesized proteins in rat hippocampal neurons.151 Recently, BONCAT and FUNCAT methods were developed in zebrafish larvae.152 It was demonstrated that Aha is metabolically incorporated into newly synthesized proteins in the larval zebrafish, in a time- and concentration-dependent manner, but with no obvious toxic effects on the organism and without influencing simple behaviors. By staining with fluorescent alkyne tags, newly synthesized proteins in whole mount larval zebrafish could be visualized by fluorescence microscopy. The identification and labeling of newly synthesized proteins by BONCAT and FUNCAT is commonly limited to proteins that contain at least one methionine. While 94% of proteinencoding genes in mammalian cells contain a methionine codon, substitution of methionine with an unnatural amino acid will operate with variable efficiency, and labeling reactions will likely proceed with variable rates.126a These factors may lead to a subset of proteins being labeled. Labeling is commonly carried out in minimal media to allow effective competition of the unnatural amino acid with the natural amino acid in the synthetase active site. Low levels of incorporation are required to avoid toxic effects that may result from proteome wide amino acid replacement, and this necessitates very sensitive detection methods. 4.3. Cell-Selective Cotranslational Labeling of Proteins

BONCAT and FUNCAT methodologies reduce sample complexity of a given proteome and permit direct analysis of the primary protein synthesis in response to stimuli. However, 4784

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Figure 26. Promoter-directed proteomic labeling with Anl (see Figure 13). (a) NLL-MetRS expression is placed under control of a promoter of interest. Only when transcriptional activity of the promoter is “on,” the NLL-MetRS is expressed and Anl is incorporated into newly synthesized proteins that can be labeled with alkyne-functionalized probes. (b) Proteomic labeling with Anl under conditions of oxidative stress. The NLLMetRS is under control of the SoxRS regulon and is activated by addition of paraquat (PQ), a superoxide-generating agent (see text for further details). Part b was adapted with permission from ref 155. Copyright 2012 American Chemical Society.

were infected with E. coli cells that constitutively expressed a plasmid-borne copy of NLL-MetRS. Anl incorporation and thereby protein labeling with an alkyne-functionalized fluorescent dye, as visualized by fluorescence microscopy, was confined to bacterial cells. Newly synthesized bacterial proteins can also be isolated from such cultures by labeling them with an enrichment tag, followed by affinity chromatography.153 This facilitated the analysis of bacterial proteins, which constitute less than 1% of the total protein content in an infection mixture. Another unnatural amino acid bearing an alkyne moiety, 2aminooctynoic acid (Aoa, Figure 13), could be incorporated residue-specifically into proteins employing the NLL-MetRS and used for cell-selective labeling.154 Aoa-modified bacterial proteins expressed in macrophage-internalized Salmonella typhimurium were labeled in a corresponding study. However, NLL-MetRS-mediated labeling employing Aoa required removal of methionine from the growth medium, while Anl incorporation is also effective in the presence of endogenous methionine.124a,125 The use of reactive unnatural amino acids that are not substrates for the endogenous translational machinery has also enabled labeling of the proteome, following transcriptional activation of a promoter driving expression of an aminoacyl-

when experiments are performed in systems that contain multiple cell types, proteins from all cell types are labeled because the specificity of the aaRS toward the unnatural amino acid is generally independent of cell identity. To achieve cellselective cotranslational labeling, it is necessary to use an unnatural amino acid that is not a good substrate for any of the endogenous aminoacyl tRNA synthetases and to provide a synthetase that will recognize this unnatural amino acid. By using an unnatural amino acid that is a poor substrate for the endogenous machinery, a lower background of protein labeling in nontargeted cells can be achieved. A mutant synthetase (NLL-MetRS, see section 3.1) that efficiently aminoacylates endogenous tRNAMet with azidonorleucine, Anl, which is not a substrate for the endogenous MetRS (Anl, Figure 13), has been identified.123 By overexpressing NLL-MetRS in certain cells, Anl incorporation can be restricted to those cells, while proteins expressed in wildtype cells use only methionine and are not labeled (Figure 25).124b This approach enables the visualization, isolation, enrichment, and identification of proteins of targeted cells in the presence of other cells. In a first demonstration of this approach, cell-selective labeling employing Anl and NLLMetRS was accomplished in a system containing mixtures of bacterial and mammalian cells. Mouse alveolar macrophages 4785

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Figure 27. Simultaneous, cell-selective metabolic labeling of two proteomes in a mixed bacterial culture is enabled by overexpressing mutant aminoacyl tRNA synthetases in distinct cells. Adapted with permission from ref 160. Copyright 2012 American Chemical Society.

tRNAMet.159 This approach was used to label, detect, and visualize newly synthesized proteins made during different stages of the cell cycle in mammalian cells. However, proteins that are subjected to cleavage of N-terminal signal sequences or proteolytic cleavage by methionyl aminopeptidases (although an N-terminal Anl attenuates cleavage activity of these enzymes) lose their Anl tag and cannot be labeled. Development of mutant mammalian synthetases that permitted incorporation of labels at internal positions in proteins would complement this approach and allow enrichment and identification also of proteins that lack N-terminal methionine residues. An engineered MetRS mutant that enabled the incorporation of the alkyne-bearing amino acid propargylglycine (Pra, Figure 13), which is not a substrate for the endogenous MetRS, was recently reported.160 This mutant MetRS, dubbed PraRS, enables incorporation of Pra at near quantitative levels into bacterial proteins expressed in a methionine auxotrophic E. coli strain. Pra-modified proteins were readily labeled with an azidemodified fluorescent dye in a Cu(I)-catalyzed click cycloaddition. The combined use of PraRS and NLL-MetRS allowed for cell-selective, differential labeling of target proteins derived from two bacterial strains cocultured in the presence of both Pra and Anl. The mixed cell-lysates were treated sequentially with a cyclooctyne-functionalized green dye (DIBO Alexa Fluor 488) to react with Anl and an azide-modified red fluorescent dye (Cy5 Azide). The in-gel fluorescence pattern enabled identification of the cellular origin of each protein (Figure 27). Recently, the construction of a bisected NLL-MetRS was reported.161 Charging of Anl to tRNAMet occurs only if both the N-terminal and the C-terminal fragments of the bisected NLLMetRS are expressed from their promoters (P1 and P2). Distinct protein labeling using either azide-modified enrichment tags or azide-bearing fluorescent dyes is apparent within 5 min after Anl has been added to bacterial cells in which both promoters have been activated. The genetic construct works as an AND gate with protein labeling as an output, which is only visible if both promoters P1 and P2 are induced. To demonstrate the approach, E. coli cells harboring this AND gate were immobilized in the center of a laminar-flow microfluidic channel, where they were exposed to various gradients of promoter inducers that drive expression of the Nterminal and C-terminal NLL-MetRS fragments and Anl. Afterward, they were fixed and treated with an alkyne-bearing fluorescent dye. The observed labeling pattern correlated with

tRNA synthetase that directs incorporation of the unnatural amino acid of interest into proteins.155 In proof of principle experiments, a gene encoding NLL-MetRS was expressed in E. coli from the PBAD promoter (Figure 26a). The patterns of protein labeling were compared under noninducing and inducing conditions following a 10 min pulse with Anl and subjecting cell lysates to alkyne-functionalized fluorescent dyes. As expected, fluorescent proteins could only be detected upon induction. The level of transcription from a promoter may reflect cellular demand for the associated proteins and may thereby reflect a cell’s physiological state. For example, transcription from the soxS promoter in E. coli drives expression of transcription factors SoxR and SoxS, which are responsible for directing expression of proteins involved in protection against damage by free radicals. Accordingly, activation of the soxS promoter is used as an indicator for oxidative stress.156 To synchronize metabolic labeling with oxidative stress response, the NLL-MetRS gene was placed under control of the SoxRS regulon by positioning it directly downstream of the soxS promoter in E. coli. This construct was then transformed into E. coli, and oxidative stress was induced by adding a superoxidegenerating agent (paraquat, PQ).157 After a 15 min pulse with Anl, tagged proteins were labeled in cell lysates by SPAAC chemistry with cyclooctyne-modified fluorophores. Evidence for tagging was only present in cells induced by addition of PQ, while proteins from cells cultured under noninducing conditions exhibited minimal background fluorescence (Figure 26b). This further demonstrates that the transcriptionally controlled expression of NLL-MetRS enables labeling of proteins expressed in response to stimuli. Expressing the mutant NLL-MetRS in mammalian cells enabled incorporation of Anl into proteins in HEK cells.158 However, Anl is not incorporated in response to all methionine codons in mammalian cells, but only in place of N-terminal methionines. This site-selective labeling (which competes with methionine incorporation and is therefore nonquantitative) is enabled by the fact that the bacterial synthetase aminoacylates mammalian initiator tRNAiMet but not elongator tRNAMet. Discrimination against mammalian elongator tRNAs is an intrinsic property of bacterial methionine tRNA synthetases: they recognize methionine tRNAs with seven-nucleotide anticodon loops (such as bacterial tRNAs and the eukaryotic initiator tRNA), but not those with nine-nucleotide anticodon loops such as the mammalian cytoplasmic elongator 4786

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Figure 28. The green fluorescent protein (GFP). (a) Structure of GFP. A magnified view of the cyclized chromophore is shown. (b) Spontaneous posttranslational modification of Ser65-Tyr66-Gly67 to produce the chromophore HBI (4-(p-hdyroxybenzylidene)-imidazolidin-5-one). (c) Expression of a genetic in-frame fusion of a protein of interest with GFP.

labeling and have had a substantial impact on biological studies, they require the use of protein fusions and/or the introduction of additional sequences into the protein of interest. This may disturb the structure and function of the protein and makes it challenging to place probes at any position in a protein. Moreover, the range of probes that can be incorporated by some of these methods is limited, and investigators have primarily focused on installing fluorescent probes. In sections 5.1.1−5.1.4, we briefly introduce four common labeling technologies and discuss their advantages and disadvantages to provide a context for a detailed discussion of site-specific protein labeling. In the subsequent sections (sections 5.2−5.3.8), we concentrate on the genetic encoding of unnatural amino acids for site-specifically installing fluorophores and probes into proteins. 5.1.1. Fluorescent Proteins. Fluorescent proteins have become indispensable tools in cell biology for monitoring molecular localization and activities of proteins and gene expression in live cells and animals.168 Their first characterized representative, the green fluorescent protein, GFP was first isolated and purified in the 1960s from the jellyfish Aequorea victoria. GFP is a ∼240 amino acid long protein (∼27 kDa) that folds into an 11-β-sheet barrel surrounding an internal distorted helix (Figure 28a).169 When exposed to blue light, GFP fluoresces green, due to a spontaneous posttranslational modification of the tripeptide Ser65-Tyr66-Gly67 in the central helix of GFP, to produce the chromophore 4-(p-hydroxybenzylidene)-imidazolidin-5-one (HBI, Figure 28b).169 HBI is nonfluorescent in the absence of the properly folded GFP scaffold, which shields it from solvent and provides a chemically complex environment for the chromophore. The hydrogen-

the strength of the individual input signals, that is, inducer concentrations.

5. SITE-SPECIFIC INCORPORATION OF UNNATURAL AMINO ACIDS FOR LABELING PROTEINS 5.1. Current Labeling Approaches

The development of techniques to image the location, structure, and dynamics of biomolecules, to image below the diffraction-limit of light, and to image single molecules would greatly benefit from approaches for site-specifically labeling proteins with small-molecule probes. The ability to rapidly label any protein at a specific position in the polypeptide with any probe of interest would provide the foundation for powerful new approaches to control and image biological function. However, the rapid, site-specific labeling of proteins in diverse contexts with user-defined probes represents an outstanding challenge for chemical biologists. There are currently four common methods used to label and image proteins within live cells.1d,e,162 First, the most common strategy involves the use of intrinsically fluorescent proteins that can be fused to the protein of interest.163 Second, selflabeling tags consisting of short peptide sequences have been developed that react directly with biarsenical dyes (tetracysteine, tetraserine tag).164 Third, a series of enzymes have been engineered that attach a fluorescently labeled molecule to one of its own amino acid residues (SNAP/Clip tag, HaloTag, and TMP tag).165 A fourth strategy involves the use of an engineered enzyme to covalently attach a labeled substrate to a tagged protein of interest (e.g., biotin ligase15a,166 and lipoic acid ligase167). While some of these approaches allow rapid 4787

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Figure 29. Self-labeling tags. (a) Labeling of proteins with the tetracysteine tag. The genetically introduced CCPGCC sequence reacts specifically with cell-permeable biarsenical-functionalized fluorescent dyes such as ReAsH. (b) Structural formulas of fluorescein and resorufin-derived biarsenical dyes FlAsH-EDT2, ReAsH-EdT2, and the rhodamine-derived bisboronic acid (RhoBo) that binds to a tetraserine motif.

bonding network and π-stacking interactions between HBI and the amino acids in the scaffold protein greatly influence the spectroscopic characteristics, like color, quantum yield, and photostablility of GFP and its various engineered variants.170 GFP and its engineered variants can be fused to any gene of interest, and translation of such a genetic in-frame fusion allows the visualization of the spatial and temporal distribution of proteins and their networks by fluorescence microscopy (Figure 28c).170,171 Since the discovery of GFP, new GFP variants with altered excitation and emission wavelengths, enhanced brightness, and improved pH resistance have been developed.172 Of particular note are recent developments for creating photoactivatable or photoswitchable fluorescent proteins, whose fluorescence can be turned on with light of a specific wavelength, or whose fluorescence excitation and emission spectra shift following illumination, respectively.173 Important representatives include Dronpa, mEos, Kaede, PAGFP, and PA-mCherry and have become indispensable tools for super resolution microscopy techniques.173a Another important and useful approach has been the engineering of GFP variants that can be used as FRET pairs, such as CFP (cyan fluorescent protein) and YFP (yellow fluorescent protein).174 GFP and its engineered variants very rarely cause phototoxicity, can be targeted to subcellular compartments, can be expressed in different tissues and also in living organisms, and can be designed to respond to a variety of biological events and signals. However, there are also some disadvantages in creating fluorescent protein fusions to visualize proteins. All GFP variants are substantial proteins of ∼27 kDa, which may perturb the structure, function, or cellular localization of the protein they are fused to. Furthermore, because of their considerable size, they can commonly only be attached to the N- or Cterminus of a protein of interest. In some instances, fluorescent proteins have been added within the sequence of a protein.175 However, this may cause significant perturbations to the overall fold, function, and dynamics of the protein of interest. In

addition, the folding and maturation of GFP and its engineered variants is far from instantaneous. The red-emitting fluorescent proteins, which are particularly useful because they can be excited independently from cellular autofluorescence, show slow maturation.176 Fluorescent protein labeling is still the method of choice for many applications. However, the photostability and photobleaching properties of small molecule fluorophores are often superior to those of fluorescent proteins. Because photostability and resistance to photobleaching are key factors for success in many imaging protocols, small molecule-based approaches should have advantages over fluorescent proteins. One alternative approach that addresses some of these limitations makes use of so-called fluoromodules, complexes formed upon the noncovalent binding of a fluorogenic dye to single-chain variable fragment (scFv) antibodies.177 The protein component, called fluorescence activating protein or FAP, binds with high affinity to the dye, leading to an up to 1000-fold enhancement of the dye’s fluorescence. FAPs can be genetically encoded and fused to a protein of interest. When a fluorogenic dye accesses the site where the FAP is expressed, the dye is bound by the FAP and fluorescence is observed. Because the binding of the dye is noncovalent, the fluorescence of the fluoromodule can also be restored after photobleaching by adding fresh dye to replace the bleached dye. Recent efforts have yielded a catalogue of fluoromodules spanning a range from blue to visible and near-IR regions of the spectrum.178 Labeling with a range of these dyes has been shown on the yeast cell surface. Labeling of intracellular proteins has not been reported so far. 5.1.2. Self-Labeling Tags. To overcome the limitations posed by the large size of fluorescent protein fusions, alternative methods for labeling proteins that employ a smaller genetically encoded tag have been developed.164a The tetracysteine tag method uses a short peptide sequence containing four cysteines (CCXXCC, where X is a natural amino acid other than cysteine) that is genetically encoded into the sequence of a 4788

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Figure 30. Self-labeling enzymes. (a) The SNAP-tag reacts with small molecules containing a benzylguanine moiety. (b) The CLIP-tag reacts specifically with O6-benzylcytosine substrates. (c) A fluorogenic version of the SNAP-tag. (d) The HaloTag forms a covalent bond between substrate and protein by removing halides from alkyl halides by nucleophilic displacement. (e) The covalent TMP-tag labeling is based on formation of a proximity driven Michael addition product between an engineered cysteine residue in the active site of EcDHFR and an acrylate-electrophile introduced into the TMP-fluorophore.

containing thiols, as does the addition (at micromolar levels) of 1,2-dithiol antidotes such as EDT or 2,3-dimercaptopropanol. Addition of millimolar concentrations of these antidotes, on the other hand, outcompetes the tetracysteine motif and makes FlAsH labeling reversible.181 In model reactions, effective dissociation constants of ≤10 pM have been established for the

target protein. This sequence, when located in a hairpin structure,179 can react specifically with cell-permeable biarsenical-functionalized fluorescent dyes such as the fluorescein derivative FlAsH and the resorufin derivative ReAsH (Figure 29a and b).180 The 1,2-ethanedithiol (EDT) derivatives of these reagents prevent the nonspecific labeling of biomolecules 4789

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however, the price paid for the gain in specificity is a larger tag size. One of the most versatile tags is the SNAP-tag, a 20 kDa mutant of the DNA repair enzyme O6-alkylguanine-DNA alkyltransferase that reacts specifically and very rapidly with small molecules containing a benzylguanine moiety, leading to covalent labeling of the SNAP-tagged protein with a synthetic probe (Figure 30a).165a Recently, a new variant of this enzyme has been developed that reacts specifically with O 6 benzylcytosine substrates, called CLIP-tag (Figure 30b).165b A fluorogenic version of the SNAP tag has been developed to reduce background fluorescence from unreacted or nonspecifically bound substrates by attaching a fluorophore and a quencher molecule to different moieties of the benzylguanine core in such a way that the fluorophore is quenched. Upon reaction with the SNAP tag, the quencher-bound guanine moiety is released, leading to a large fluorescence increase of the fluorophore, which remains attached to the protein tag (Figure 30c).194 The fluorogenic SNAP tag was combined with a fast-labeling variant of the SNAP-tag (SNAPf), which shows a 10-fold increase in reactivity toward benzylguanine substrates. This approach enabled the wash-free, real-time visualization and dynamics of SNAP-tagged proteins in cell lysates and living cells.194,195 Both SNAP-tag and CLIP-tag fusion proteins have been used for live-cell imaging with a variety of different synthetic fluorescent probes. Importantly, SNAP- and CLIP-tags are orthogonal and can be used together in one cell to label two proteins with two different fluorescent substrates. SNAP- and CLIP-tag fusion proteins have been used for live-cell imaging with a variety of different synthetic fluorescent probes.165b A major advantage of these tags is the great variety of different synthetic fluorescent probes that can be used as substrates, and this has been important in developing and improving superresolution techniques like STED (stimulated emission depletion) microscopy and STORM (stochastic optical reconstruction microscopy).196 Apart from SNAP- and CLIP-tag, the bacterial enzyme haloalkane dehalogenase has been engineered as a self-labeling protein tag (HaloTag).165c Haloalkane dehalogenase removes halides from alkyl halides by nucleophilic displacement. An engineered variant of this enzyme that has been mutated in the active site traps the covalent ester intermediate between the alkyl residue of the alkyl halide and the enzyme, enabling the covalent attachment of synthetic labeled substrates (Figure 30d). Similar to SNAP- and CLIP-tags, genetic in-frame fusions of the HaloTag have been used to investigate the localization and trafficking of proteins in living cells.197 Another strategy for labeling proteins in living cells exploits the high-affinity interaction between E. coli dihydrofolate reductase (EcDHFR) and folate analogues like trimethoprim (TMP).165d When EcDHFR is fused to a protein of interest, the tagged protein can be labeled with high affinity (KD ≈ 1 nM) and selectivity (KD for mammalian DHFRs > 1 μM)198 with TMP-based fluorescent probes.165d The TMP-tag has been used to label proteins with high efficiency and selectivity both in vitro and inside live cells.1e,199 Recently, a covalent version of the TMP-tag has been developed by installing a unique cysteine residue on EcDHFR in a position to react with an acrylamide electrophile added to the TMP-probe via a classic proximityinduced Michael addition, allowing a more permanent label for advanced applications as well as lower concentrations of labeling reagent (Figure 30e).200 This initial design of a covalent TMP-tag, however, suffered from slow in vitro

optimized binding sequence CCPGCC with association kinetics of >105 M−1 s−1.181 This sequence has been further improved to yield two high-affinity 12 amino acid long biarsenical binding motifs (FLNCCPGCCMEP and HRWCCPGCCKTF) for labeling in mammalian cells.182 An advantage of the tetracysteine motif results from the fact that biarsenical probes are highly fluorogenic: they are essentially nonfluorescent in solution and undergo a dramatic enhancement in fluorescence upon binding to the protein, which enhances the signal-to-noise ratio in labeling reactions.181 The biarsenical dyes are, however, not without problems. Concerns include the nonspecific hydrophobic interaction of these dyes with membranes or off-target cellular proteins, which necessitates washing steps in the presence of competing thiols, as well as the overall cellular toxicity of the ligands, and the effect of the tag on the protein function and localization. Nevertheless, the tetracysteine reporter group has been used to image a variety of proteins in vivo including some whose localization and function were known to be perturbed by GFP labeling.183 Live cell applications include studies of mRNA translation,184 amyloid185 and microtubule formation,182 transport of viruses,186 and receptor activation.187 Furthermore, extensions of the methodology have been developed for electron microscopy imaging,181,188 affinity chromatography,189 western blotting,189b and in vivo pulse labeling experiments.188 A system named bipartite tetracysteine display uses the tetracysteine motif to study protein interactions and folding.190 In this approach, the two halves of a tetracysteine motif (i.e., a pair of cysteines) are introduced into recombinant protein domains distal in primary sequence but proximal in the folded or assembled state, and addition of the biarsenical dyes allows studying protein−protein interactions and protein folding. This system has been used to reveal conformational changes in EGFR activation upon ligand binding at the mammalian cell surface.191 To extend the range of fluorophores that can be targeted to tetracysteine tags, a modular approach has been adopted, wherein the bis-arsenical targeting group is separated from the fluorophore through a spirolactam tether (SplAsH-tag). A fluorophore of choice can then be added to the tether. This modularity offers high flexibility; however, it comes at the cost of reduced fluorogenicity and cell-permeability.192 To circumvent the cytotoxicity related to biarsenical dyes, a rhodamine-derived bisboronic acid (RhoBo, Figure 29b) that had previously been described as a monosaccharide sensor was developed for protein labeling.193 RhoBo binds with even higher affinity to a tetraserine motif (SSPGSS) introduced into a protein of interest than to monosaccharides. Similar to FlAsH and ReAsH, RhoBo is cell permeable and fluorogenic, but because it does not contain arsenic, it is less toxic to live cells and it does not suffer from nonspecific background fluorescence arising from thiol exchange. However, while the CCPGCC motif is absent from the human proteome, some endogenous proteins contain SSPGSS-containing sequences, which may lead to some off-target labeling in live cells.164b 5.1.3. Self-Labeling Enzymes. The success of self-labeling tags has inspired the development of related methods for covalently attaching small molecules to proteins. Many of these approaches rely on the fusion of an engineered enzyme to the protein of interest. The enzyme then covalently attaches a labeled probe to the protein. Self-labeling proteins are expected to provide higher labeling specificity than self-labeling tags; 4790

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Figure 31. Enzyme-mediated labeling. (a) General scheme for enzyme-mediated labeling: a recognition sequence (peptide tag) is fused to the protein of interest. An engineered enzyme recognizes the peptide sequence and covalently attaches a labeled substrate to it. (b) Phosphopantetheinyl-transferases (PPTAses) transfer a CoA-derivative to a small carrier protein (CP) fused to a protein of interest (POI). (c) Biotin ligase (BirA) specifically ligates a biotin derivative to a lysine residue within an acceptor peptide (AP) sequence fused to the POI; the ketonemodified biotin is then reacted with a hydrazide labeled probe. (d) Engineered variants of lipoic acid ligase (LplA) site-specifically modify an acceptor peptide (AP), fused to the POI, with an azide-modifed short chain fatty acid substrate, which is subsequently labeled via SPAAC. (d) Cterminal labeling of proteins using Sortase.

cysteine residue installed outside of the TMP-binding pocket.202 While self-labeling proteins like the tags described above allow the exploitation of the wider repertoire of spectrochemical properties represented by synthetic organic fluorophores, an additional step of washing out the free dye is often required to ensure a low fluorescent background. Furthermore, some of these tags are relatively large proteins with sizes similar to GFP. Thus, related issues as for fluorescent proteins apply: the fused

kinetics, limiting its use in live cell imaging. By optimizing the positioning of the cysteine residue as well as structural features of the acrylamide electrophile, a second generation covalent TMP-tag with a more rapid labeling half-life was recently developed, offering a robust and rapid reagent for live cell imaging.201 Recently, a fluorogenic version of the TMP-tag was reported, in which a TMP-fluorophore-quencher molecule was synthesized with the quencher attached to a leaving group, which upon TMP-binding to EcDHFR would be cleaved by a 4791

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5.2. Incorporation of Fluorescent Unnatural Amino Acids

tag might hamper or hinder the structure, function, or localization of the protein of interest. 5.1.4. Enzyme-Mediated Labeling. An alternative labeling method for proteins makes use of an engineered enzyme that recognizes a peptide sequence or small protein fused to a protein of interest and covalently attaches a labeled substrate to it. Ideally, this approach should combine the small tag size provided by self-labeling tags with the high specificity conferred by enzymes (Figure 31a).1d A well-established approach uses the bacterial phosphopantetheinyl-transferases (PPTases) AcpS203 (from E. coli) or Sfp204 (from B. subtilis) to transfer a CoA-derivative to a small carrier protein (9 kDa) fused to a protein of interest (Figure 31b).205 These enzymes do not recognize the corresponding carrier proteins in mammalian cells and can therefore be used orthogonally within a mammalian system. The labeling is, however, restricted to the cell surface as the CoA-bearing labeling reagent would interfere with intracellular processes involving CoA. Efforts in evolving and engineering the carrier protein have enabled a reduction of the tag size from 9 kDa to an 8−12 amino acid long tag.206 Biotin ligase has also been used to label specific proteins. E. coli biotin ligase (BirA) specifically biotinylates a lysine residue within a 15 amino acid long peptide tag, which is orthogonal to the peptide sequence recognized by mammalian biotin ligases. Consequently, mammalian proteins fused to the BirA recognition sequence peptide can be specifically modified with biotin and visualized with streptavidin-fluorophore conjugates. BirA also accepts a biotin isostere containing a keto group, which can be exploited for labeling with hydrazide or aminooxy compounds (Figure 31c).15a Biotin ligases from other species can also accept alkyne and azide-modified biotin substrates, enabling labeling via Staudinger ligation or CuAAC (see sections 2.2 and 2.3).166 BirA can be expressed within mammalian cells and used for intracellular labeling. Limitations and off-target reactivity are however represented by endogenous biotinylated proteins. Recently engineered versions of lipoic acid ligase (LplA) have been employed to site-specifically modify proteins with modified short chain fatty acid substrates (Figure 31d).167 These engineered enzymes are able to efficiently transfer substrates containing reactive bromides, azides, alkynes, aldehydes, hydrazides, trans-cyclooctenes, as well as photocross-linkers and the fluorescent dye coumarin to proteins tagged with the 13-amino acid long LplA acceptor peptide (LAP) recognition sequence.15b,95c,207 This approach has been used to label specifically tagged proteins on the cell surface, intracellularly, and in the nucleus. Furthermore, lipoic acid ligase-mediated labeling can be used in parallel with biotin ligase to label two independent proteins in the same cell. However, every newly labeled substrate requires engineering of the lipoic acid ligase active site because the substrate has to fit into the active cavity within the enzyme. Another method uses sortase (SrtA) to mediate specific protein labeling. The enzyme recognizes a 5 amino acid long peptide sequence near the C-terminus of its target site, cleaves a specific peptide bond within this sequence, and forms an amide bond between the new C-terminal amino acid and an Nterminal glycine of a polyglycine species (Figure 31e). The method has been used for both C-terminal208 and Nterminal209 labeling of cell-surface proteins.

The site-specific, genetically encoded incorporation of fluorescent amino acids would allow the generation of fluorescent proteins. A couple of fluorescent amino acids have been incorporated site-specifically into proteins using amber suppression. The fluorescent amino acid L-(7-hydroxycoumarin-4-yl)ethylglycine (HceG, Figure 32) has been encoded in E.

Figure 32. Structural formulas of fluorescent unnatural amino acids HceG, DansA, and Anap.

coli using an evolved MjTyrRS/tRNACUA pair and incorporated site-specifically into sperm whale myoglobin.210 Incorporation was confirmed by SDS-PAGE and ESI−mass spectrometric analyses. The coumarin fluorophore is sensitive to solvent polarity, and the site-specifically incorporated HceG was used to monitor unfolding of myoglobin in the presence of guanidinium chloride. Furthermore, HceG was incorporated site-specifically into enhanced cyan fluorescent protein (eCFP) to create a fluorescent protein with an increased Stokes shift.211 Site-specific incorporation of HceG into the bacterial tubulin homologue FtsZ in E. coli was used to visualize in vivo assembly of this cytoskeletal protein into a contractile midcell ring during cytokinesis. HceG-modified FtsZ retained its functionality, while fusion of FtsZ to a fluorescent protein like GFP impaired its function.212 An environmentally sensitive dansyl amino acid (DansA, Figure 32) was also genetically encoded and site-specifically incorporated into proteins expressed in S. cerevisiae using an engineered E. coli leucyl-tRNA synthetase/tRNACUA pair. DansA was incorporated into human superoxide dismutase and used as an environmentally sensitive reporter of protein unfolding.213 A fluorescent unnatural amino acid, based on the environmentally sensitive fluorophore 6-propionyl-2-(N,N-dimethyl)aminonaphthalene (Anap, Figure 32), has been synthesized and incorporated into proteins expressed in S. cerevisiae and in mammalian cells using an engineered variant of the E. coli leucyl-tRNA synthetase/tRNACUA pair.127b,214 Anap-modified proteins were used to study ligand-induced local conformational changes in proteins and biomolecular interactions in vitro and to localize proteins in living mammalian cells. With respect to established methods for fluorescently labeling proteins, the incorporation of fluorescent unnatural amino acids introduces minimal perturbations to protein structure and is therefore unlikely to interfere with the function and localization of proteins as approaches using fusions or tagging can. However, the amino acids that can currently be encoded use quite short wavelengths, making them nonideal for in vivo imaging. In addition, it may be challenging to wash out the background of free fluorescent amino acid (which is added to the cells at millimolar concentrations) to give a high signal4792

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Figure 33. Labeling proteins via incorporation of unnatural amino acids that can be chemoselectively labeled. (a) An unnatural amino acid bearing a unique bioorthogonal functionality is introduced site-specifically into a protein via genetic code expansion and then chemoselectively labeled with an externally added chemical probe. (b) Rate constants of chemoselective reactions for which one of the partners can be genetically encoded in form of an unnatural amino acid. (Note that the reaction rate of CuAAC is dependent on Cu(I) concentrations used in the labeling reaction. Rate constants of 10−200 M−1 s−1 are achieved using Cu(I) coordinating ligands and copper concentration of 100−500 μM as employed in cell-surface labeling experiments in mammalian cells; see section 2.3.)

Figure 34. Structural formulas of unnatural amino acid useful for chemoselective labeling that have been incorporated site-specifically into proteins via genetic code expansion.

5.3. Incorporation of Unnatural Amino Acids with Bioorthogonal Handles

to-noise ratio, a problem not encountered when using

A flexible approach to protein labeling is presented by the sitespecific installation of bioorthogonal chemical groups into a

fluorescent proteins. 4793

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5.3.2. Incorporation of Azides. The reaction of azides with terminal alkynes through a copper(I)-catalyzed [3 + 2] cycloaddition reaction (see section 2.3) has been extensively used for the in vitro labeling of biomolecules. p-Azidophenylalanine (p-AzF, Figure 34) has been incorporated into proteins in E. coli and into peptides displayed on phage using an evolved MjTyrRS/tRNACUA pair.219 Furthermore, p-AzF has been genetically incorporated into proteins in eukaryotic cells using an evolved bacterial TyrRS/tRNACUA pair.14c,45a,215c,d Purified p-AzF containing proteins have been labeled in vitro with fluorescent dye- or PEG-derivatized alkynes in copper-catalyzed CuAAC reactions. Proteins containing p-azidophenylalanine can also be labeled in a Staudinger ligation (section 2.2) with fluorophore tethered phosphines.38a,219c Similarly, Nε-(oazidobenzyloxycarbonyl)-L-lysine (o-AzbK, Figure 34) has been incorporated into glutathione transferase (GST) using an engineered PylRS/tRNA pair and labeled via Staudinger ligation with a fluorescein-triarylphosphine derivative.38b The PylRS/tRNACUA pair was also used to genetically incorporate aliphatic azides Nε-(2-azidoethoxy)carbonyl-L-lysine (AzK, Figure 34) into proteins in E. coli.220 In contrast to the aromatic azides previously incorporated, this aliphatic azide is easier to use, because it is photostable and can be synthesized in a few simple steps. Another aliphatic azide, an azide-bearing cyclic pyrrolysine analogue (ACPK, Figure 34), has been synthesized and genetically incorporated into proteins in E. coli and mammalian cells using an engineered version of the PylRS/ tRNACUA pair.221 While azides are commonly considered to be stable in cells, azides introduced into proteins via genetic code expansion are often reduced to the corresponding amines, making labeling at azides nonideal. 5.3.3. Incorporation of Alkynes. Several alkynes can be genetically incorporated into proteins, and these can be labeled in bioorthogonal CuAAC reactions with azide containing probes. p-Propargyloxy-L-phenylalanine (PrgF, Figure 34) has been genetically incorporated into proteins in E. coli using an evolved MjTyrRS/tRNACUA pair222 and into yeast using an evolved bacterial TyrRS/tRNACUA pair.45a,215c,d Mutant proteins were purified and labeled in vitro with fluorescent dye- or PEG-derivatized azides. A pyrrolysine analogue tagged with an acetylene group Nε-(3-ethynyltetrahydrofuran-2carbonyl)-L-lysine (EtcK, Figure 34), requiring multistep synthesis, was incorporated into proteins in E. coli using the PylRS/tRNACUA pair, and proteins were modified selectively with an azido coumarin dye by CuAAC chemistry.223 Furthermore, for FRET experiments, double labeling of EtcK with azido coumarin was performed in conjunction with cysteine labeling with an Alexa488 C5-maleimide dye on calmodulin. The PylRS/tRNACUA pair directs the incorporation of a simple aliphatic alkyne (AlkK, Figure 34) that can be synthesized in high yield in a few simple steps.220 The mutant protein was quantitatively labeled with azide-modified cyanine dyes and biotin as characterized by in-gel fluorescence, western blot, and mass spectrometry. Recently, another lysine derivative bearing a terminal alkyne (AlkK2, Figure 34) has been incorporated site-specifically into bacterial proteins using an evolved PylRS/tRNACUA pair. Tagged proteins were labeled with iodide derivatives in a Sonogashira coupling.86 The strain-promoted azide−alkyne cycloaddition (SPAAC, section 2.3) has also been used to chemoselectively modify proteins.27 The genetic encoding of cyclooctyne bearing lysines

protein by genetic code expansion and the subsequent labeling of these proteins via specific chemoselective reactions. This provides an emerging route to site-specific protein labeling with tailor-made biophysical probes (Figure 33a).107 As mentioned in section 2, the labeling reaction between the unnatural amino acid bearing the bioorthogonal handle and the externally added chemical probe has to proceed under biologically compatible conditions, in which the reactants form a product with each other but do not cross-react with the many other chemical functional groups found in proteins or other biomolecules in living cells and organisms.1a,b This approach to labeling proteins is more flexible than the direct genetic introduction of the probes, and allows labeling of proteins with essentially any sort of probe including a range of fluorophores, crosslinking agents, and cytotoxic molecules. Many of the functional groups that can be encoded for chemical protein labeling methods participate in reactions that have rate constants on the order of 10−4−1 M−1 s−1, which are 8 to 4 orders of magnitude slower than enzymatic labeling approaches (Figure 33b) and have precluded their use in many in vivo applications. In section 5.3.8, however, we will discuss recent advances in the development of very rapid chemoselective reactions, including the incredibly rapid reaction between tetrazines and strained alkenes and strained alkynes, and the encoding of the corresponding reaction partners, which allow very rapid site-specific labeling of cellular proteins within living cells. 5.3.1. Incorporation of Ketones. As described in section 2.1, ketones react with hydrazines and hydroxylamines. The keto-containing amino acids p-acetylphenylalanine (p-AcF, Figure 34),14b,c,215 m-acetylphenylalanine (m-AcF, Figure 34), 1 9 and p-benzoylphenylalanine (p-BzF, Figure 34)14c,215a,c,d,216 have been incorporated in E. coli using the MjTyrRS/tRNACUA pair and in eukaryotic cells using bacterial Tyrosyl-tRNA synthetase/tRNA pairs. The introduction of these unnatural amino acids into recombinant proteins has allowed their site-specific labeling with hydrazine- or hydroxylamine-modified fluorescent dyes and EPR probes in vitro. Furthermore, the MjTyrRS was engineered to incorporate a βdiketone-containing phenylalanine (dkF, Figure 34) in E. coli. DkF-modified proteins were labeled using a hydroxylamine containing dye.217 Double site-specific protein labeling via a genetically encoded ketone and a cysteine has allowed FRET probes to be introduced into proteins for single molecule FRET spectroscopy.215b Both p-AcF and p-BzF were incorporated into G-protein coupled receptors (GPCRs) and labeled with fluorescent probes in vitro. Apart from reaction with hydrazines and hydroxylamines, p-BzF also provides the possibility of photocross-linking to identify precise sites of protein−protein interaction.215a More recently, an aliphatic keto-containing amino acid (2-amino-8-oxononanoic acid, Aon, Figure 34) has been incorporated site-specifically into proteins in E. coli, using an evolved PylRS/tRNA pair. Here, a modified protein was labeled with a biotinylated alkoxyamine derivative and a ketoreactive hydrazide dye.218 As discussed in section 2.1, the reaction of ketones with hydroxylamines or hydrazines requires an acidic pH, which is not ideal for some proteins. Furthermore, the reaction is quite slow, exemplified by second-order rate constants in the range of 10−4 M−1 s−1, thereby requiring both high concentrations of labeling reagent and long reaction times, which precludes its use for in vivo labeling applications. 4794

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Figure 35. Palladium-catalyzed cross-coupling reactions on proteins. (a) Mizoroki− Heck reaction between p-iodophenylalanine and a labeled vinyl reagent. (b) Sonogashira reaction between p-iodophenylalanine and a labeled terminal alkyne. (c) Suzuki cross-coupling reaction between a genetically encoded p-boronophenylalanine and a bodipy aryliodide probe.

(CoK1 and CoK2, Figure 34) into proteins in E. coli using an engineered PylRS/tRNACUA pair has been achieved.224 Modified proteins were labeled in live E. coli cells via incubation at 37 °C for 3−6 h with an azide coumarin dye.97,224 A lysine derivative bearing a more strained cyclooctyne, bicyclo[6.1.0]nonyne (BCNK, Figure 34), has also been incorporated into proteins in E. coli and mammalian cells using engineered PylRS/tRNACUA pairs.95a,97 Incorporated BCNK reacted considerably faster than CoK1 in strainpromoted click cycloadditions with an azide coumarin dye.97 The real advantage of BCNK, however, lies in its exquisite reactivity with tetrazine probes in inverse-electron-demand Diels−Alder cycloadditions as described in section 5.3.8.95a 5.3.4. Incorporation of 1,2-Aminothiols. Variants of the orthogonal PylRS/tRNACUA pair have been discovered that direct the efficient, site-specific incorporation of Nε-L-thiaprolylε L-lysine (ThiPK, Figure 34), N -D-cysteinyl-L-lysine (DCysK, 225 ε Figure 34), and N -L-cysteinyl-L-lysine (LCysK, Figure 34) into proteins in E. coli.226 These proteins, modified with a unique 1,2-aminothiol group, were efficiently and specifically labeled with a cyanobenzothiazole-modified dye (see section 2.7). Furthermore, in combination with cysteine-maleimide labeling, this approach allowed the dual labeling of proteins with distinct probes at two distinct, genetically defined sites.226 The 1,2-aminothiol-CBT condensation reaction shows faster kinetics (k2 ≈ 9 M−1 s−1)74b than many other chemoselective reactions for which one of the partners can be genetically encoded. However, the reaction is limited to in vitro applications, because the 1,2-aminothiol functionality forms adducts with pyruvate and possibly other metabolites in vivo, and deprotection is required prior to labeling. 5.3.5. Incorporation of Reaction Partners for PdCatalyzed Cross-Coupling. Palladium-mediated cross-coup-

ling reactions are among the most powerful reactions in organic chemistry for constructing new carbon−carbon bonds.227 Palladium is absent from all known biological systems, and the reaction partners of a Suzuki coupling, arylhalides and boronic acids, do not occur naturally in polypeptides, either as posttranslational modifications or as cofactors. p-Iodo-Lphenylalanine (p-IF, Figure 34) containing Ras protein was prepared using an E. coli cell free translation system.81a This protein was conjugated in vitro to a biotin-tethered vinyl via the Mizoroki−Heck reaction, using a water-soluble Pd-TPPTS (triphenylphosphine-3,3′,3″-trisulfonate) catalyst. However, after 50 h of incubation at 5 °C, only 2% of biotinylated product could be detected by mass spectrometry (Figure 35a). Reaction of a p-IF-modified protein with a biotinylated alkyne in a Sonogashira reaction using Pd(OAc)2, TPPTS, and CuOTf afforded 25% of coupled product after 80 min incubation at 6 °C (Figure 35b).81b A variant of the MjTyrRS/tRNACUA pair has been evolved to genetically encode p-IF into proteins in E. coli, and an engineered version of the bacterial Tryrosyl-tRNA synthetase has been created to incorporate p-IF into proteins in eukaryotic cells.14c,215c,d,228 p-Borono-L-phenylalanine (p-BorF, Figure 34) has been incorporated into proteins in E. coli using an engineered variant of the MjTyrRS/tRNACUA pair, and a p-BorF-modified Zdomain-protein was coupled to a BODIPY aryliodide probe in a Suzuki cross-coupling reaction using a Pd0-dibenzylidene acetone (pd-DBA) catalyst in 30% yield at 70 °C (Figure 35c).81c All of these initial reports on palladium-catalyzed reactions suffered from very low yields and/or required high temperatures, making them inadequate for quantitative chemoselective labeling under biologically relevant conditions. 4795

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Figure 36. Photoclick chemistry on proteins. (a) Reaction of 2-(p-methoxyphenyl)-5-phenyltetrazole with O-allyl phenyl ether. (b) Tetrazolemodified proteins can be labeled in vitro with a fluorescein-tethered fumarate using 254 nm UV irradiation. (c and d) Model reactions of ethyl-1methylcycloprop-2-enecarboxylate with two different tetrazoles proceed with second-order rate constants of 60 and 5 M−1 s−1, respectively. (e) Twophoton-triggered 1,3-dipolar cycloaddition between naphthalene-based tetrazoles and alkenes.

was observed after 1 h at 37 °C with millimolar concentrations

Recently, a new water-soluble palladium catalyst was reported for protein labeling: the complex Pd(OAc)2(ADHP)2 (see Figure 10, section 2.9).82 Using this catalyst, the in vivo labeling of a bacterial cell-surface protein, which had been sitespecifically modified by genetically encoding p-IF, with a fluorescent boronic acid was demonstrated.83b Clear labeling

of both palladium catalyst and boronic acid. No toxic effects on E. coli could be observed at these concentrations. While these observations are encouraging, additional experiments will be required to investigate the cell permeability of this and future 4796

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Figure 37. Site-specific protein labeling via anilines. (a) Aniline reacts under oxidative conditions with phenylene diamino groups to yield imines. (b) Aniline-modified proteins are conjugated to peptides containing N-terminal phenylene diamine groups.

constant dropped by more than 10-fold (k2 ≈ 5 M−1 s−1, Figure 36d). Incubation of CpK-modified myoglobin with 2-(pmethoxyphenyl)-5-phenyltetrazole and photoirradiation at 302 nm for a period of 1−12 min revealed formation of the fluorescent pyrazoline adduct by in-gel fluorescence analysis, with the highest fluorescent intensity reached after 10 min. Mass spectrometric analysis confirmed greater than 85% conversion of the CpK-labeled myoglobin to the pyrazoline product. To examine whether CpK can direct bioorthogonal labeling in mammalian cells, HEK 293 cells were transfected with a plasmid encoding for the CpK synthetase as well as with a plasmid encoding for eGFP containing an amber codon. Cells were grown in the presence of 4 mM CpK, treated with 40 μM of photoreactive tetrazole for 1.5 h followed by photoirradiation at 365 nm for 2 min, and finally imaged using a fluorescence microscope. Green fluorescence, arising from eGFP expression, colocalized with pyrazoline fluorescence; however, not all green fluorescent cells were labeled, which may indicate some variability in the cell-permeability of the tetrazole. As compared to other genetically encoded, chemoselective labeling reactions, the main advantage of photoclick chemistry lies in its potential for spatiotemporally controlled protein labeling in mammalian cells, which, however, will require the development of new highly reactive tetrazole reagents that can be laser-activated at wavelengths that are less harmful to living cells.73c,d Recently, the design and application of naphthalene-based tetrazoles that can be activated by two-photon excitation with a 700 nm femtosecond pulsed laser has been reported. This enables photoclick chemistry with acrylamide groups to be performed in living organisms with improved spatiotemporal control.230 Upon photoirradiation with a femtosecond pulsed laser, the appropriately functionalized tetrazoles form a highly reactive 1,3-dipole that reacts with a suitable dipolarophile such as acrylamide in a fluorogenic 1,3-dipolar cycloaddition to yield fluorescent pyrazolines (Figure 36e). An alkene-modified protein was prepared by genetically encoding Nε-acryloyl-Llysine (AcrK, Figure 34)231 and labeled site-specifically in vitro by photoirradiating a naphthalene-modified tetrazole. Moreover, microtubules were labeled in live cells by incubating and photoirradiating cells with a fumarate-modified microtubule binding drug and the appropriately modified tetrazole. 5.3.7. Incorporation of Anilines. In addition to the use of alkynes, azides, ketones, terminal alkenes, and tetrazoles as

Pd catalysts for intracellular labeling, especially in mammalian cells. Recently, the incorporation of two new unnatural amino acids, bearing either iodo or alkyne functionalities (AlkK2, pIpK, see Figure 34), into proteins in Gram-negative bacterial cells has been reported, creating proteins with handles for palladium-mediated cross-coupling reactions.86 A reaction screening system led to the discovery of Pd(NO3)2 as a simple and highly efficient and biocompatible reagent for fluorescent labeling of target proteins via a copper-free Sonogashira coupling reaction in vitro and in living E. coli cells. 5.3.6. Incorporation of Photoclick Reaction Partners. Components of the photoclick reaction (see section 2.6, Figure 7) have been genetically encoded, and site-specific protein labeling via the reaction has been demonstrated. O-Allyltyrosine (O-AllY, Figure 34) has been incorporated site-specifically into proteins in E. coli using an engineered variant of the MjTyrRS.229 O-AllY-modified proteins have been labeled with a variety of diaryltetrazole probes under UV irradiation at 302 or 365 nm.71 The formation of the products was confirmed by in-gel fluorescence and mass spectrometric analyses. Kinetic analysis revealed that reaction of 2-(p-methoxyphenyl)-5phenyltetrazole with O-allyl phenyl ether proceeded with a rate constant of 0.95 M−1 s−1 (Figure 36a).71b The photoclick reaction was also performed in live E. coli cells expressing OAllY-containing Z-domain proteins and followed by fluorescence microscopy. O-AllY-containing Z-domain proteins showed a rapid fluorescence increase after the bacteria were illuminated at 302 nm for less than 1 min without the need for additional incubation.71a In addition, the genetic incorporation of p-(2-tetrazole)phenylalanine (p-TzF, Figure 34) into proteins in E. coli by an evolved MjTyrRS variant was reported.73a p-TzF-modified myoglobin was labeled in vitro with a fluorescein-tethered fumarate using 254 nm UVirradiation (Figure 36b). Recently, the site-specific incorporation of a 3,3-disubstituted cyclopropene-containing lysine (CpK, Figure 34) into proteins in E. coli and mammalian cells using an engineered PylRS variant has been reported.73b The cycloaddition reaction of 2-(p-methoxyphenyl)-5-phenyltetrazole with model substrate ethyl-1-methylcycloprop-2-enecarboxylate showed a second-order rate constant of ∼60 M−1 s−1 (Figure 36c). Using a water-soluble, more biocompatible, 365 nm wavelength photoreactive tetrazole, however, the rate 4797

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Figure 38. Inverse-electron-demand Diels−Alder cycloadditions between tetrazines and dienophiles for labeling proteins. (a) Fluorescent labeling of tetrazine-modified proteins with strained-trans-cyclooctene probes. (b) Structural formulas of tetrazine probes discussed in this Review. (c) Structural formulas of tetrazine-reactive dienophiles discussed in this Review.

Recently, aaRS/tRNA pairs for the efficient site-specific incorporation of several unnatural amino acids bearing components of the very rapid inverse-electron-demand Diels−Alder cycloaddition were reported. A tetrazine-containing phenylalanine derivative104 (TetF, Figure 34) was introduced at a specific site in a protein expressed in E. coli via an engineered variant of the MjTyrRS. TetF-modified GFP was then derivatized with a strained trans-cyclooctene, and reaction rates were determined both in vitro and in E. coli to be considerably faster than many other site-specific labeling reactions (880 and 330 M−1 s−1, respectively). Furthermore, Tet-F-modified GFP was fluorescently labeled with a strained trans-cyclooctene-bearing diacetyl-fluorescein dye (Figure 38a). Strained alkene and alkyne functionalities, including norbornenes 96a,b (NorK, Figure 34), bicyclo[6.1.0]nonynes95a,97 (BCNK, Figure 34), trans-cyclooctenes95a,d (TCOK, Figure 34), and 1,3-disubstituted cyclopropenes (Elliot, T., Chin, J. W., unpublished results), have also been encoded. By engineering PylRS/tRNACUA pairs for these unnatural amino acids, it was possible to incorporate them site-specifically into proteins expressed in E. coli and mammalian cells. A series of unsymmetrical aryl-tetrazines that contained a unique reactive group for functionalization with biophysical probes were developed that showed different reactivities toward dienophiles according to their substitution patterns with electrophilic moieties (tet1, tet2, tet3, tet4, and tet5, Figure 38b).96a Coupling these tetrazines to certain red fluorophores led to a substantial decrease in fluorescence with respect to the fluorescence of the parental fluorophores due to an energy transfer to the tetrazine chromophore, which absorbs

chemical handles for protein labeling, proteins can also be sitespecifically labeled via an aniline. Francis and co-workers developed an oxidative coupling strategy, which involves the modification of aniline-containing proteins under oxidative conditions in aqueous solution (Figure 37a).232 They further demonstrated the utility of the reaction by site-specifically labeling proteins via genetically encoding the unnatural amino acid p-aminophenylalanine (p-AmF, Figure 34).233 A viral capsid was modified by genetically encoding p-AmF into a coat protein and conjugated to peptides containing N-terminal phenylene diamine groups via this strategy (Figure 37b).233a Furthermore, p-AmF-modified viral capsids were conjugated to nucleic acid aptamers via this oxidative coupling strategy, enabling the capsids to preferentially bind to targeted cells.234 This coupling strategy uses the oxidant sodium periodate, which can oxidize cysteine and methionine side chains in the proteins; furthermore, commonly used reductants such as DTT (Dithiothreitol) or TCEP (Tris(2-carboxyethyl)phosphine) that are commonly present in buffers will inhibit the reaction.233a 5.3.8. Incorporation of Tetrazines and Dienophiles for Inverse-Electron-Demand Diels−Alder Reactions. Many of the chemoselective reactions discussed so far, for which one of the partners can be genetically encoded, show modest coupling rates (10−4−1 M−1 s−1), which makes it challenging to use them for site-specific protein labeling in vivo. Genetically encoded cyanobenzothiazole condensations226 and photoclick reactions73b have been developed with rate constants reaching 10−60 M−1 s−1, but commonly have side reactions in vivo or require short-wavelength light for activation. 4798

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Figure 39. Spectroscopic and kinetic analysis of the tetrazine-dienophile reaction. (a) Structural formulae of tetrazine-fluorophore conjugate T-tet1 (i.e., tetrazine tet1 (see Figure 38b), coupled to a red emitting tetramethylrhodamine (TAMRA) fluorophore shown in red) and T-tet1 after reaction with Nor (see Figure 38c, shown in blue). The fluorescence of T-tet1 is turned on upon cycloaddition with Nor, leading to a 5−10-fold gain in fluorescence intensity. (b) Kinetic analysis of the Diels−Alder reaction. The rate constants were determined under pseudo first-order conditions with an excess of dienophile over tetrazine by following the exponential decay of UV absorbance at 320 nm over time. From the observed rate constants (k′), obtained at different concentrations of dienophiles, the second-order rate constants of the individual reactions were determined. The table shows the second-order rate constants for different combinations of tetrazine and dienophile (for structural formulas, see Figure 38).

between 510 and 550 nm.235 Importantly, however, the fluorescence of these tetrazine-dye conjugates is turned on upon cycloaddition with the corresponding strained alkenes or alkynes (Nor, BCN, TCO, and sTCO, Figure 38c), leading to a 5−10-fold gain in fluorescence intensity (Figure 39a). The “turn-on” fluorescence upon cycloaddition may be a major advantage for site-specific protein labeling where probe wash-

out is not desired or possible. The rate constants of different dienophiles (Nor, BCN, TCO, see Figure 38c) with various tetrazines (tet1, tet2, te3, tet4, see Figure 38b) were determined under pseudo first-order conditions using stopped flow techniques, by following the exponential decrease in the UV absorbance of the tetrazines.95a,96a The second-order rate constants of these reactions approach 105 M−1s−1, which is 4799

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Figure 40. Specific and ultrarapid labeling of extracellular and intracellular proteins in live mammalian cells. (a) Site-specific labeling of epidermal growth factor receptor (EGFR)-GFP fusion on the cell surface. In the presence of BCNK or TCOK (for structures, see Figure 33), HEK 293 cells produced full-length EGFR-GFP that can be visualized at the cell membrane by fluorescence microscopy. To demonstrate the specific labeling of EGFR-GFP that contained BCNK or TCOK with tetrazine fluorophores, cells were treated with the tetramethylrhodamine (TAMRA)-tetrazine conjugate T-tet1 (for structural formula see Figure 39), washed, and the red fluorescence arising from tetramethylrhodamine labeling as well as the green fluorescence arising from expression of full-length EGFR-GFP were imaged. Clear labeling of cells that contained EGFR-BCNK-GFP or EGFR-TCOK-GFP was observed within 2 min, and tetramethylrhodamine fluorescence clearly colocalized with cell-surface EGFR-GFP fluorescence. No labeling was observed for cells in the same sample that did not express EGFR-GFP, and cells bearing EGFR-BocK-GFP (BocK = Nε-tert-butyloxycarbonyl-L-lysine) were not labeled with T-tet1. (b) Specific and rapid labeling of a nuclear protein in live mammalian cells. A transcription factor, jun, with a C-terminal mCherry fusion was expressed from a gene bearing an amber codon in the linker between JunB (jun) and mCherry. In the presence of BCNK and the corresponding synthetase/tRNA pair, jun-BCNK-mCherry protein was produced in HEK cells and, as expected, localized in the nuclei of cells. Labeling with the cell-permeable conjugate CFDA-tet5 (see Figure 38, CFDA = diacetyl-fluorescein) resulted in green fluorescence that colocalized with the mCherry signal at the first time point analyzed (after 15 min of labeling and 90 min of washing). No specific labeling was observed in nontransfected cells in the same sample or in control cells expressing jun-BocK-mCherry, further confirming the specificity of intracellular labeling.95a

initially weakly fluorescent, become strongly fluorescent once attached to the protein of interest via the chemical reaction, making the signal-to-noise of this labeling approach superior. This approach may be extended to site-specific protein labeling in animals and represents a step-change in approaches to labeling proteins. We foresee that this will have a broad impact on labeling and imaging studies. While isomerization of the trans double bond in TCOK has not been problematic in labeling to date, the stability of this bond in long-term labeling studies and diverse biological environments will need to be investigated. Moreover, while it has been demonstrated that a site-specifically incorporated

extraordinarily fast and approaches the rate of many enzymemediated labeling procedures (Figure 39b). Proteins with site-specifically incorporated dienophile-bearing unnatural amino acids were labeled selectively and very rapidly with tetrazine fluorophores in E. coli and in live mammalian cells both in vitro and in vivo. With this approach, it was possible to label extracellular as well as intracellular proteins at genetically encoded unnatural amino acids via a chemoselective reaction within live mammalian cells (Figure 40a and b). These labeling reactions use low (nanomolar) concentrations of labeling reagents, are very specific, are very rapid, and many of the red tetrazine fluorophores, which are 4800

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Notes

NorK, BCNK, and TCOK can be specifically labeled in a target protein expressed at low levels without significant labeling of the rest of the E. coli proteome,95a,96a the specificity of the reaction still merits a more detailed investigation.

The authors declare no competing financial interest. Biographies

6. CONCLUSIONS AND FUTURE CHALLENGES In summary, a range of chemoselective reactions have been used to label isolated biomolecules, cell surface biomolecules, and intracellular biomolecules at physiological temperatures and pressures. Many of these reactions proceed under aqueous conditions and produce nontoxic or no byproducts. The rates of these chemoselective reactions span 9 orders of magnitude (see Figure 33b), and the recent development of rapid reactions promises applications of labeling to previously inaccessible biological problems. In many cases, a deeper investigation of the chemoselectivity of these reactions with respect to potentially competing reactions with cellular molecules present at high abundance will advance our understanding of the scope and limitations of these reactions for studying biology. Efforts to quantitatively profile the specificity of reactions with respect to classes of molecules in cells, for example, proteomes,95a,96a will be particularly helpful in understanding how to improve the specificity of reactions further. Established bioorthogonal reactions have been used to label and in some cases enrich proteomes synthesized at defined times and in mixtures of cells. These approaches are providing new biological insights, and new opportunities are likely to emerge by combining new, rapid, chemoselective chemistry with cotranslational proteome labeling. The development of reactions that are chemoselective and rapid under biologically relevant conditions is being rapidly translated into approaches for selective protein labeling in cells and animals via genetic code expansion. Encouragingly for in vivo imaging, although genetic code expansion approaches commonly direct unnatural amino acid incorporation in response to the amber codon, there appears to be minimal background labeling resulting from incorporation and labeling at endogenous amber codons in E. coli, the only system where this has been investigated.95a,96a This will need to be investigated more quantitatively and in a wider range of biological systems to determine whether very low abundance proteins can be selectively imaged and whether initial observations extend to additional cell types. Additional challenges for intracellular imaging include the wash-out of the labeling probe, but emerging approaches to cell permeable and turn-on probes may help address this challenge.235,236 The approaches developed for site-specific protein labeling are poised to have a deep impact on the imaging and control of protein dynamics, structure, and function in living cells and animals.

Kathrin Lang received her Ph.D. degree from the University of Innsbruck, Austria, in 2008 under the supervision of Prof. Ronald Micura. Since then she has been working as a Postdoctoral Fellow and Investigator Scientist at the Medical Research Council Laboratory of Molecular Biology (MRC LMB) in Cambridge, UK, in the research groups of Dr. Venki Ramakrishnan and Prof. Dr. Jason Chin. In the Chin lab she has developed approaches for the rapid, site-specific labeling of proteins in and on live cells via genetic code expansion. Her general research interests lie in the interdisciplinary area of chemistry and biology, applying concepts from organic chemistry to develop new tools to study fundamental biological questions. At the start of 2014 she was appointed as a Mössbauer Professor (Assistant Professor) in Synthetic Biochemistry at the Technische Universität München, Institute for Advanced Study, Germany.

Jason Chin is currently a Programme Leader at the Medical Research Council Laboratory of Molecular Biology (MRC LMB) in Cambridge, UK, where he is also Head of the Centre for Chemical & Synthetic Biology (CCSB). He is Professor of Chemistry & Chemical Biology at the University of Cambridge, and holds a joint appointment at the University of Cambridge Department of Chemistry. He is also a fellow in Natural Sciences at Trinity College, Cambridge.

AUTHOR INFORMATION Corresponding Authors

*E-mail: [email protected]. *E-mail: [email protected].

REFERENCES (1) (a) Sletten, E. M.; Bertozzi, C. R. Angew. Chem., Int. Ed. 2009, 48, 6974. (b) Lim, R. K.; Lin, Q. Chem. Commun. (Cambridge, U. K.) 2010, 46, 1589. (c) Prescher, J. A.; Bertozzi, C. R. Nat. Chem. Biol. 2005, 1, 13. (d) Hinner, M. J.; Johnsson, K. Curr. Opin. Biotechnol. 2010, 21, 766. (e) Jing, C.; Cornish, V. W. Acc. Chem. Res. 2011, 44, 784.

Present Address †

Department of Chemistry, Technische Universität München, Institute for Advanced Study, Lichtenbergstrasse 4, 85748 Garching, Germany 4801

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dx.doi.org/10.1021/cr400355w | Chem. Rev. 2014, 114, 4764−4806