Selective chemical labeling of proteins

Abstract

Over the years, there have been remarkable efforts in the development of selective protein labeling strategies. In this review, we deliver a comprehensive overview of the currently available bioorthogonal and chemoselective reactions. The ability to introduce bioorthogonal handles to proteins is essential to carry out bioorthogonal reactions for protein labeling in living systems. We therefore summarize the techniques that allow for site-specific “installation” of bioorthogonal handles into proteins. We also highlight the biological applications that have been achieved by selective chemical labeling of proteins.

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Xi Chen

Dr. Xi Chen was born in Sicuan, China in 1986. He received his MSc in Chemistry at the National University of Singapore and PhD degree (2012) in Protein Chemistry at Ulm University. Since 2012, he conducted his postdoctoral research under the supervision of Dr. Yao-Wen Wu at the Max-Planck Institute for Molecular Physiology and the Chemical Genomics Centre of the Max-Planck Society. His research focuses on chemical protein labeling and the development of photochemical approaches for controlling protein function in live cells.

Yao-Wen Wu

Dr. Yao-Wen Wu was born in Jiangxi, China in 1980. He received his BSc in Chemistry from Sun Yat-sen University in 2001 and his MSc in Organic Chemistry from Tsinghua University in 2004. After obtaining Dr rer. nat. (summa cum laude) in 2008 at TU Dortmund working at the Max Planck Institute of Molecular Physiology in Dortmund, he then conducted his postdoctoral research in cell biology at King's College London. Since 2010 he has been leader of an Otto Hahn group at the Max Planck Institute in Dortmund. Since 2012 he has been group leader of the Chemical Genomics Centre of the Max Planck Society. His research interests include molecular mechanism of membrane trafficking and autophagy, protein chemical modification, chemical probes for visualizing cellular events, chemical and optogenetics.

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1. Introduction

Selective chemical labeling of proteins is highly useful for probing and controlling protein functions in vitro and in living systems.^1–10 Since proteins carry many copies of different functional groups, it is demanding to selectively label a protein at a defined site. Additionally, the labeling conditions have to be mild and reactions should be able to undergo in aqueous solution. Moreover, the labeling reagents should also be orthogonal to other functionalities in living systems when a protein is to be labeled in vivo.^11,12 These restrictions make it really challenging to label proteins in a chemoselective and site-specific manner.

Recent years have seen tremendous progress in chemical protein labeling. There are already some excellent reviews on bioorthogonal reactions and selective chemical labeling of proteins.^{2–4,6,12–18} In this review, we compare the features of reported bioorthogonal reactions and chemical tagging approaches, e.g. the reaction rate or time, reaction conditions, reagents, etc. This could be helpful to identify the most suitable labeling reaction for a particular application. Moreover, we highlight the applications of established chemical labeling techniques to tackle biological problems.

2. The toolbox of bioorthogonal chemistry for protein labeling

Bioorthogonal reactions are robust and invaluable tools for chemical protein labeling.^4,6 Typically, selective protein labeling is accomplished by incorporation of bioorthogonal groups into a protein, followed by chemoselective modifications.¹⁸ This approach is also designated as “tag-and-modify”.¹⁹ A variety of bioorthogonal reactions have been developed, which can be classified into: (1) condensation reactions through carbonyls, (2) “click” reactions through azides, (3) inverse electron-demand Diels–Alder cycloadditions (DA_INV) and other cycloaddition reactions, (4) transition metal-catalyzed coupling and decaging reactions, and (5) labeling reactions at cysteine residues (Table 1). In parallel, many elegant approaches have been established to selectively equip proteins with bioorthogonal handles (discussed in later sections), fulfilling the requirements for the subsequent bioorthogonal modification.

Table 1 The toolbox of bioorthogonal chemistry and chemoselective reactions

Reactant A	Reactant B	Product	k (M^−1 s^−1) or t	Features	Reference(s)
t: reaction time. (A) RT: room temperature; ABAO: aminobenzamidoxime; KAHA: α-ketoacid-hydroxylamine; ACL: aldehyde capture ligation. (B) PBS: phosphate buffered saline; TBTA: tris[(1-benzyl-1H-1,2,3-triazol-4-yl)methyl]amine; CuAAC: copper-catalyzed azide alkyne cycloaddition; SPAAC: strain-promoted azide alkyne cycloaddition; BCN: bicyclononyne. (C) TCO: trans-cyclooctene; ED: electron-donating. (D) TQ-ligation: thiovinyl ether o-quinolinone quinone methide ligation; oQQM: o-quinolinone quinone methide; EW: electron-withdrawing. (E) BIAN: bis-imine of acenaphthenequinone and mesitylamine; dba: dibenzylidene acetone. (F) TCEP: tris(2-carboxyethyl)phosphine; Dha: dehydroalanine; MSH: O-mesitylenesulfonylhydroxylamine; CBT: cyanobenzothiazole; NCL: native chemical ligation; MESNA: 2-mercaptoethanesulfonate; MPAA: 4-mercaptophenylacetic acid.
A: Condensation through carbonyls
			0.006 (pH 7, RT)	Oxime ligation: p-phenylenediamine (p-PDA) or aniline, etc. as a catalyst	Dirksen (2006)^20
			0.29 (pH 7, RT, 10 mM p-PDA)		Wendeler (2014)^21
			PBS (pH 6.5, RT) 0.0087, or 0.46 (1 mM 5MA)	Hydrazone ligation: aniline or 5-methoxyanthranilic acid (5MA), etc. as a catalyst; ∼85% yield in 5 h	Crisalli (2013)^22
			18 h (pH 6.5, 37 °C)	Pictet-Spengler reaction: formation of a stable C–C bond; suitable for N-terminal protein labeling	Sasaki (2008)^23
			10 (pD 4.5)	Pictet-Spengler ligation: ligation is faster under lower pH conditions; formation of a stable C–C bond	Agarwal (2013)^24
			2 (pD 6)
			4.17 (pH 6)	Hydrazino-Pictet-Spengler ligation: ligation proceeds fastest under pH 6; formation of a stable C–C bond	Agarwal (2013)^25
			40 (pH 4.5)	ABAO ligation: formation of a fluorescent dihydroquinazoline (λmax 490 nm); quantitative labeling in 1 h	Kitov (2014)^26
			1.61 (pH 7.4, RT, 10 μM CuSO4, air)	Under air; requires Cu(II) or Zn(II) catalyst; quantitative labeling in 2 h	Ji (2014)^27
			24 h (pH 7, RT)	Mukaiyama-Adol condensation: 11–55% conversion; stable	Alam (2010)^28
			4 h (≥10% H2O in DMSO, 60 °C)	KAHA ligation: formation of a native amide bond; 56–76% yield	Bode (2006)^29
					Pattabiraman (2012)^30
			1.65 (DMF, RT); 40 h [10% DMF in PBS (pH 7)]	ACL: selective N-terminal protein labeling is possible at pH 7; ∼70% labeling of ubiquitin	Raj (2015)^31
B: Click reactions through azides
			0.003 (PBS)	Staudinger-Bertozzi ligation: phosphine is prone to oxidation	Saxon (2000)^32
			0.0074 (H2O)	Traceless Staudinger ligation: needs transient protection of phosphine by BH3; 80% yield at pH ≥ 7.5	Nilsson (2001)^33
					Tam (2007)^34
			6–24 h (H2O, RT)	Staudinger-phosphite ligation: phosphite is stable against air oxidation; up to 90% yield	Serwa (2009)^35
			∼3 [50 μM Cu(I) + 50 μM TBTA, DMSO : H2O 1 : 9]	CuAAC: ligand for copper catalyst is required; up to 94% yield	Rostovtsev (2002)^36
					Presolski (2010)^37
			up to 0.0024 (H2O, 25 °C)	Oxanorbornadiene cycloaddition: metal-free, but is slow	van Berkel (2007)^38
			0.0024–0.076	SPAAC: cyclooctynes are susceptible to thiols	Agard (2004)^39
					Baskin (2007)^40
			0.057–0.96	SPAAC: cyclooctynes are lipophilic; nonspecifically stick to serum proteins and cellular membranes	Ning (2008)^41
					Jewett (2010)^42
			0.29; 2–2.9 (with electron-deficient azide)	SPAAC: BCN is readily accessible; the reaction rate is accelerated using an electron-deficient azide	Dommerholt (2010/2014)^43,44
C: Inverse electron-demand Diels–Alder cycloadditions (DA INV )
			210–∼8.6 × 10^5 (PBS, 37 °C)	TCO is lipophilic and isomerizes over time; 100% conversion in 5 min	Blackman (2008)^45
			437–2672	BCN is readily accessible; almost quantitative conversion	Lang (2012)^46
			0.12–9.46	Norbornene is stable	Devaraj (2008)^47
			0.03–27	Cyclopropene is small and stable; ED groups increase reactivity; quantitative labeling	Patterson (2012)^48
					Elliott (2014)^49
			0.39 (DMSO : H2O 12 : 88, 20 °C)	Used for affinity based protein profiling	Engelsma (2014)^50
			0.017–0.041	Terminal olefin only reacts with highly reactive tetrazines	Niederwieser (2013)^51
			0.12–0.58 (THF : H2O 1 : 1)	Tertiary isonitrile is required; the reaction is slow	Stöckmann (2011)^52
D: Other dipolar cycloadditions
			39 (MeCN : H2O 1 : 9, 25 °C)	Strain-promoted alkyne-nitrone cycloaddition (SPANC): some nitrones are prone to hydrolysis; up to 95% yield	Ning (2010)^53
			0.043 (CDCl3, RT)	Diazo-strained alkyne cycloaddition: the reaction rates are similar to the equivalent reactions of azide	McGrath (2012)^54
					Josa-Culleré (2014)^55
			0.054 (21 °C, 55% MeOH in H2O)	Strain-promoted sydnone-BCN cycloaddition: quantitative reaction in organic solvent and in aqueous buffer	Wallace (2014)^56
			0.25 (PBS/EtOH)	Quadricylane ligation: quadricyclane is stable; the product is light sensitive	Sletten (2011)^57
			N.A.	Nitrile oxide-norbornene cycloaddition: nitrile oxide may cross react with nucleophiles; mostly used for labeling of nucleic acids	Gutsmiedl (2009)^58
			0.0015–0.028 (37 °C, 1 : 5 MeCN : H2O)	TQ ligation: oQQM generated in situ; EW-groups on oQQM accelerate the reaction	Li/Dong (2013)^59
					Zhang (2015)^60
			0.79 (MeCN : PBS 1 : 1)	Tetrazole-alkene photo-click reaction: irradiation wavelength is tetrazole dependent; nitrile imine generated in situ; up to quantitative yield	Wang (2008/2009)^61,62
			2 min (PBS, RT): give 41% labeling of lysozyme	Azirine ligation: azirine generates a reactive nitrile ylide intermediate in situ	Lim (2010)^63
E: Transition metal-catalyzed coupling/decaging reactions
			30 min (ligand, pH8, 37 °C)	Suzuki–Miyaura coupling: boronic acids are moderately toxic; >95% conversion	Chalker (2009)^64
			30 min (Cu ion, ligand, RT)	Sonogashira coupling: Mg(II) inhibits the binding of Pd(II) to the protein	Kodama (2007)^65
			2–5 h (pH 8, 30% tBuOH/H2O, RT to 37 °C)	Olefin cross metathesis: requires ruthenium catalyst; up to 90% yield	Lin (2008)^66
			24 h (Pd(OAc)2, BIAN, O2, pH 7, RT)	Aqueous oxidative Heck reaction: reaction opens to air; up to full conversion if excessive catalyst and boronic acid are used	Ourailidou (2014)^67
	allyl2Pd2Cl2 or Pd(dba)2		180 min (10 μM Pd(dba)2, 37 °C)	Useful for in vivo protein activation	Li (2014)^68
F: Labeling at cysteine residue(s)
			<30 min (PBS, RT)	TCEP should be added to maintain cysteine resides in a reduced form	Kim (2008)^69
			5 min (pH 7.5, RT, 5 : 95 MeCN : H2O)	X = I, Br, Cl, OTf; quantitative labeling. Cy = cyclohexyl-	Vinogradova (2015)^70
			90 min (pH 9.0, 4 °C or RT)	Dha can be derived from cysteine by MSH (pH8.0, 4 °C, 20 min); >95% labeling	Bernardes (2008)^71
			9.19 (PBS)	CBT ligation: side reactions with thiols of CBT	Ren (2009)^72
			pH 7.0	NCL: thiol catalysts are used, e.g., MESNA, MPAA	Dawson (1994)^73
			After reduction of disulfide: 2 h (pH 7.8, 4 °C)	S–S intercalation: reaction under mildly basic pH; formation of a stable product	Shaunak (2006)^74
			TCEP, 1 h; then pH 8, RT, 3 h	Acetone crosslinking: the introduced ketone allows oxime ligation; cross linking also stabilizes helical structures	Assem (2015)^75

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2.1. Condensation through carbonyls

Ketones and aldehydes can react with hydroxyamine and hydrazide compounds under aqueous conditions to form stable oxime and hydrazone linkages, respectively. Oxime ligation⁷⁶ is slow under neutral pH conditions and therefore an aniline catalyst is required.²⁰ Recently, an improved catalyst p-phenylenediamine (p-PDA) was reported, which displays a higher water solubility and a better catalytic efficacy (10–120 times faster) than aniline.²¹ With respect to the condensation using hydrazide (hydrazone ligation), electronic and acid/base effects strongly influence the reaction efficiency at pH 7.4. For instance, carbonyl compounds with neighboring acid/base groups (e.g. carboxylate) form hydrazones at accelerated rates of up to 2–20 M⁻¹ s⁻¹.⁷⁷ In addition, in the presence of a 5-methoxyanthranilic acid (5MA) catalyst, the condensation rate can be substantially enhanced 84-fold (6.6 M⁻¹ s⁻¹, 1 mM 5MA) compared to the reaction without a catalyst (0.08 M⁻¹ s⁻¹) at pH 6.5 (Table 1A).²²

Aside from oxime ligation and hydrazone ligation, aldehydes and ketones can undergo a Pictet-Spengler reaction with β-arylethylamines,⁷⁸ which has been used for protein labeling at the N-terminus.²³ To date, several modified versions have been introduced, including Pictet-Spengler ligation²⁴ and hydrazino-Pictet-Spengler ligation.²⁵ Additionally, proteins engineered with an N-terminal aldehyde tag can be labeled via the Mukaiyama-Adol condensation using silyl ketene reagents with the formation of a stable C–C bond.²⁸ KAHA (α-ketoacid-hydroxylamine) ligation allows the condensation between an α-ketoacid and hydroxylamine or 5-oxaproline to form a native amide bond. This ligation has been a valuable alternative to native chemical ligation (NCL)⁷³ to join two unprotected peptide fragments in peptide synthesis.^29,30 Owing to the rapid association between aldehydes and amines, an aldehyde has been elegantly employed as an amine-capture auxiliary in aldehyde capture ligation (ACL). In ACL, a C-terminal selenobenzaldehyde ester can interact with the N-terminus of a peptide/protein. A native amide bond linkage is formed following a Se→N acyl shift. The ACL has been used for site-specific N-terminal modification of ubiquitin (Table 1A).³¹

A recently introduced reaction is ABAO (2-aminobenzamidoxime) ligation. ABAO combines an aniline moiety for iminium-based activation of the aldehyde with a nucleophilic group at the ortho-position to the amine for intramolecular ring closure. In addition to the rapid condensation reaction kinetics (up to 40 M⁻¹ s⁻¹), the condensation forms a fluorescent dihydroquinazoline derivative, making it possible to develop fluorogenic aldehyde-reactive probes.²⁶ Alkyl aldehydes can also efficiently couple with aryl diamines under mild conditions (RT, neutral aqueous solution) in the presence of Cu(II) or Zn(II) ions via an oxidative condensation process.²⁷ This reaction forms stable benzimidazole linkages and has been utilized to label the T4 lysozyme protein with an aldehyde dye (Table 1A).

With respect to labeling biomolecules in live cells or organisms, carbonyl compound-related condensations have not been widely used. This is because the catalysts are usually toxic, and endogenous ketones and aldehydes, e.g. glucose and pyruvate, would interfere with the labeling reaction. Nevertheless, ketones and aldehydes are generally not present on the cell surface. Therefore, carbonyls serve as useful chemical handles for labeling biomolecules on the cell surface using hydrazide or aminooxy probes.^20,79–83

2.2. “Click” through azides

Azide is a small and stable group, which has a unique dipole for a variety of bioorthogonal reactions.⁸⁴ Click reactions using azides include Staudinger ligation,³² traceless Staudinger ligation,^33,34 Staudinger-phosphite ligation,³⁵ copper-catalyzed azide alkyne cycloaddition (CuAAC),³⁶ strain-promoted azide alkyne cycloaddition (SPAAC) and oxanorbornadiene cycloaddition (Table 1B).³⁸ Among these reactions, CuAAC and SPAAC appear to be the most popular and are discussed in detail in this section.⁴

CuAAC is a hallmark of bioorthogonal chemistry that was reported independently by Sharpless³⁶ and Meldal⁸⁵ in 2002. Its application as a bioorthogonal reaction revolutionized our ability to modify and manipulate proteins.^86,87 CuAAC becomes popular mainly due to the following reasons: (1) the azide and alkyne groups are highly specific toward each other but remain inert to other chemically active molecules in live systems; (2) the reaction produces a regioselective 1,4-triazole product which is stable and inert; (3) CuAAC exhibits fast reaction kinetics (∼3 M⁻¹ s⁻¹ in the presence of 50 μM Cu(I) and 50 μM TBTA)³⁷ and various ligands have been developed to stabilize Cu(I) and further increase the reaction speed.

Cu ions catalyze the production of reactive oxygen species, leading to cytotoxicity.⁸⁸ This limits the application of CuAAC in living systems. By choosing an appropriate ligand, CuAAC can be biocompatible with minimal cytotoxicity while showing an increased reaction rate.⁸⁹ A panel of these ligands is summarized in Scheme 1A. Copper-chelating azides bring the Cu(I) ion into close proximity and thereby significantly increase the reaction rate (Scheme 1B).⁹⁰ The ligand BTTP (3-[4-({bis[(1-tert-butyl-1H-1,2,3-triazol-4-yl)methyl]amino}methyl)-1H-1,2,3-triazol-1-yl]propanol) stabilizes the Cu(I) ion while speeding up the click reaction (Scheme 1A). In addition, its complex with Cu(I) is cell permeable and non-cytotoxic, facilitating CuAAC in live E. coli cells.⁹¹ Using this reaction, an environment-sensitive fluorogenic fluorophore (alky-4DMN) was site-specifically introduced into HdeA in both the periplasm and cytoplasm of E. coli. HdeA, an acid-stress chaperone that adopts pH-dependent conformational changes, was genetically encoded with an azide-carrying unnatural amino acid ACPK at residue 58 within its pH-responsive region. The resulting hybrid pH indicator enables compartment-specific pH measurement to determine the pH gradient across the E. coli cytoplasmic membrane (Scheme 1C).⁹¹

Scheme 1 Typical CuAAC ligands (A) and variants of improved CuAAC (B and C).

Fluorogenic azide probes display substantial fluorescence enhancement upon cycloaddition reaction and therefore confer labeling with minimal background (Fig. 1A). Based on photo-induced electron transfer (PET) mechanism, highly fluorogenic blue-emissive azidomethyl substituted anthracene (A) and its analogues have been generated.⁹² Substitution at the 3 or 7 position of coumarin has a strong impact on its fluorescence properties. Guided by this principle, 3-azido substituted coumarins (B) show an 80-fold increase in fluorescence intensity upon the cycloaddition reaction.⁹³ The quantum yield (Φ, QY) of a probe after a click reaction can be calculated by density function theory (DPF), allowing rational design of fluorogenic azide probes.⁹⁴ Using this approach, the green-emission azido-fluorescein (C) has been designed, which exhibits a fluorescence enhancement of 29 to 34-fold upon cycloaddition with different alkynes.⁹⁵ Fluorogenic green- to far red-emitting CalFluors enable sensitive detection of biomolecules under no-wash conditions (D, E, F).⁹⁶

Fig. 1 (A) A panel of highly fluorogenic azide probes including a selection of CalFluors; the maximal excitation/emission wavelengths refer to the value after the click reaction; R = –(CH₂)₂O(CH₂)₂N⁺Me₂(CH₂)₃SO₃⁻. (B) Target identification using an affinity-based probe (AfBP) by CuAAC.

An important application of CuAAC lies in the target identification of biologically active small molecules in biomedical research and drug discovery.⁹⁷ A bioactive compound is typically derivatized with a photo-crosslinking moiety (e.g. diazirine) and a terminal alkyne, which is denoted as an affinity-based probe (AfBP) (Fig. 1B). By photo-crosslinking, the AfBP probe in situ captures the protein target and forms a stable protein–ligand complex in the cell. Subsequent labeling of the target proteins using azide probes is followed by separation via gel-electrophoresis and determination via mass spectroscopy.⁹⁷ Among many bioorthogonal groups, a terminal alkyne is a well suited tag as it is small with minimal interference of protein–ligand interactions, chemically inert, and can be easily modified via CuAAC.

Although the cytotoxicity of Cu(I) can be reduced by using ligands, copper-free click chemistry is more straightforward. Strain-promoted azide–alkyne cycloaddition (SPAAC) and other copper-free click reactions³⁸ without the requirement for a metal catalyst have been developed (Table 2A). The first SPAAC reaction for protein labeling was reported by Bertozzi and coworkers in 2004 using a strained cyclooctyne.³⁹ However, the reaction rate is slow (k = 2.4 × 10⁻³ M⁻¹ s⁻¹), which is comparable to Staudinger ligation (k = 3 × 10⁻³ M⁻¹ s⁻¹). To date, new variants of strained alkynes have been developed with improved properties, such as enhanced specificity, reduced lipophilicity⁹⁸ and increased reaction rates (Table 2B).

Table 2 Variants of strained alkynes for SPAAC

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There are several ways to improve strained alkynes.⁹⁹ The first strategy is to modulate the electronic properties by introducing electron-withdrawing (EW) groups, e.g. fluorine, near the triple bond.¹⁰⁰ Examples include MOFO¹⁰¹ and DIFO.⁴⁰ The second approach is to fuse the cyclooctynes with rigid aromatic rings, leading to an enhancement of reactivity by increasing ring strain. The ring-fused cyclooctynes, including DIBO,^41,102 DIBAC,¹⁰³ COMBO¹⁰⁴ and BARAC,⁴² show a 25–400-fold increase in the reaction rate.¹⁰⁵ However, synthesis of these cyclooctynes is usually laborious. Bicyclononyne (BCN), a cyclopropane-fused cyclooctyne, shows relatively fast reaction kinetics (0.3–1 M⁻¹ s⁻¹) toward azides and can be facilely prepared in only three steps.⁴³ Using an electron-deficient azide, e.g. 4-azido-1-methylpyridin-1-ium iodide, the cycloaddition reaction rate with BCN can be further increased to 2–2.9 M⁻¹ s⁻¹.⁴⁴ By introducing both EW groups and ring strain, difluorobenzocyclooctyne (DIFBO) shows only a moderate increase of reactivity (0.22 M⁻¹ s⁻¹) with a significant reduction in stability.¹⁰⁶ The third strategy is to shorten the ring size as exemplified by the 7-membered tetramethylthiazacycloheptyne (TMTH),¹⁰⁷ cyclohexyne and cyclopentyne.¹⁰⁸ However, these compounds have poor stability (Table 2B). The copper-free click reactions have been used for antibody-free western blot analysis,¹⁰⁹ visualization of glycosylation on cell surfaces¹¹⁰ and protein labeling inside live cells.¹¹¹

2.3. Inverse electron-demand Diels–Alder cycloaddition (DA_INV) and other cycloadditions

The inverse electron-demand Diels–Alder cycloaddition (DA_INV) occurs between an electron-rich dienophile (e.g. strained alkene/alkyne) and an electron-poor diene, typically 1,2,4,5-tetrazine (Table 3A). DA_INV represents so far the most rapid bioorthogonal reaction (up to 8.6 × 10⁵ M⁻¹ s⁻¹).^45,112 In contrast, the reaction rate of SPAAC is only up to 1 M⁻¹ s⁻¹. Theoretical calculation suggests that the rapid reaction rate can be attributed to the much higher interaction energy between tetrazine and trans-cyclooctene (TCO) in comparison to the interaction energy between an azide and a strained alkyne.¹¹³

Table 3 Reactivity of strained alkenes and tetrazines

a Protein labeling reaction. b The reaction kinetics values with BCN which is roughly 10–15 times smaller than with TCO⁴⁶. c Reaction with sTCO; stability (%) refers to the intact tetrazine in fetal bovine serum (FBS) at 37 °C after 10 h.¹¹⁵

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Tetrazines are highly reactive so that they can also readily react with strained alkynes, isonitriles⁵² and even terminal alkenes. Among these reactions, the reaction between tetrazine and BCN or cyclopropene has proven to be the most useful. Tetrazine reacts with BCN in a rapid fashion with a reaction rate constant of 3000 M⁻¹ s⁻¹.⁴⁶ In comparison to TCO that requires complicated synthetic procedures, BCN is readily available through organic synthesis. Compared to BCN and TCO groups, cyclopropene (Cpp) is much smaller, and therefore has been employed as a “minimalist” tag for live-cell imaging and affinity-based protein labeling.¹¹⁴

Alkenes and alkynes display different reactivity toward tetrazine (Table 3B).¹¹⁶ sTCO features a cyclopropane ring, which brings additional strain and increases the reaction rate of the cycloaddition up to 160-fold in comparison to TCO.¹¹⁶ The carbamate bond near the trans-double bond in TCO* reduces the chance of nucleophilic attack. Thus, TCO* shows better stability than TCO with only 2-fold reduction in reactivity toward tetrazine.¹¹⁷ The reaction rate of BCN with tetrazine is about 10 times slower than that of TCO.⁴⁶ Norbornene is ca. 10 000 times less reactive than TCO, as it is more bulky and exhibits less ring strain than TCO.^46,47 Other alkenes, including Cpp,^48,49,114 acylazetine⁵⁰ and terminal alkene,⁵¹ are less reactive, and therefore can react only with highly reactive tetrazines.

Different tetrazines show varied reactivity toward strained alkenes (Table 3C). Studies have been conducted to evaluate the reactivity and stability of tetrazines.¹¹⁵ The introduction of EW substituents can substantially increase the reaction rate. However, it is a double-edged sword. Increase in the reactivity of tetrazine often results in the reduction of stability and lifetime in the serum. ED substituents on the other hand decrease the reactivity of tetrazine.¹¹⁵ Besides electronic effects, steric effects also play a crucial role. In general, there is a trade-off between reactivity and stability. The rate constants of tetrazine reactions with TCO as well as their stability are summarized in Table 3C.

Green- and red-emitting fluorophores display electronic interactions with tetrazine chromophores that have absorption maxima at 500–525 nm. As a consequence, tetrazine-conjugated fluorophores often show reduced fluorescence. After a cycloaddition reaction, tetrazine is deconjugated and loses its quenching capability. Hence, many tetrazine-fluorophores feature fluorogenic properties, such as BODIPY-FL, Oregon Green 488, BODIPY-TMR, VT680,¹¹⁸ TAMRA, and fluorescein.¹¹⁹ These probes show a moderate turn-on ratio (up to 20-fold) (Fig. 2A). By adapting through-bond energy transfer (TBET) for fluorescence quenching, Weissleder and coworkers developed green-emitting BODIPY-tetrazine probes with up to 1600-fold turn-on ratio¹²⁰ and blue-emitting coumarin-tetrazine probes with up to 11 000-fold fluorescence enhancement.¹²¹ The tetrazine moiety is attached at the meta-position to the fluorophore on a rigid phenyl ring (Fig. 2A). Under these conditions, the tetrazine group is perpendicular to the fluorophore moiety, leading to the collinear alignment of two dipoles.¹²¹

Fig. 2 (A) Fluorogenic tetrazine-fluorophores; (B) dual protein labeling using orthogonal DA_INV pairs for super-resolution microscopy (SRM).

To achieve dual- or multi-labeling of proteins, mutually orthogonal reactions are desirable.¹²² By fine-tuning the DA_INV reactions, “selectivity enhanced” DA_INV reactions have been applied to sequential dual-color labeling of insulin receptors (IRs) and virus-like particles (VLPs) on the cell surface of HEK293T cells, facilitating dual-color super-resolution microscopy (Fig. 2B).¹¹⁷

In addition to CuAAC, SPAAC and DA_INV, which are popularly used for protein labeling, other dipolar cycloaddition reactions are summarized in Table 1D, including strain-promoted alkyne-nitrone cycloaddition (SPANC),⁵³ diazo-strained alkyne cycloaddition,^54,55 nitrile oxide-norbornene cycloaddition,⁵⁸ quadricylane ligation,⁵⁷ TQ-ligation,^59,60 strain-promoted sydnone-BCN cycloaddition,⁵⁶ and plenty of photo-triggered cycloaddition reactions, such as the tetrazole-alkene photo-click reaction^61,62 and azirine ligation.⁶³

2.4. Transition metal-catalyzed couplings and decaging reactions

Biocompatible transition metal-catalyzed couplings include the Suzuki–Miyaura coupling,⁶⁴ Sonogashira coupling, and olefin metathesis. These reactions can be performed under mild and aqueous conditions, despite the necessity to use a transition metal catalyst (Table 1E).¹⁵ Suzuki–Miyaura coupling requires an iodophenyl group, a water soluble palladium (0) catalyst and an appropriate ligand. Recently, an improved ligand, 1,1-dimethylguanidine, has been employed for aqueous Suzuki–Miyaura coupling.¹²³ Suzuki–Miyaura coupling has been successfully used for protein glycosylation and labeling at the cell surface.¹²⁴ Sonogashira coupling is another Pd-catalyzed coupling reaction.⁶⁵ An alkyne group is incorporated into the protein and subsequently reacts with an iodophenyl probe. Sonogashira coupling has been used for the modification of ubiquitin in live cells.^11,15,125

Terminal olefins can undergo an oxidative Heck reaction with boronic acid in the presence of Pd(OAc)₂/BIAN catalysts. This reaction has been used for the site-specific labeling of 4-oxalocrotonate tautomerase (4-OT).⁶⁷ Olefin metathesis is the redistribution of fragments of alkenes by scission and regeneration of carbon–carbon double bonds.^117,118 Hence, in order to avoid undesired cross coupling products, a terminal olefin and a more reactive thiovinyl ether are required.¹²⁶ Water soluble ruthenium catalysts have been developed to mediate the reaction in aqueous solution.^126,127

Chemical protein labeling not only makes proteins visible but also renders them controllable. Recently, palladium catalysts have been used to manipulate protein function in cells. Pd-mediated cleavage of the propargyl carbamate group leads to the generation of a free lysine residue. The protected lysine analogue can be genetically and site-specifically incorporated into a protein using an unnatural amino acid (UAA) mutagenesis technique (discussed in the later section). This strategy enables protein activation in living cells by decaging the lysine residue located at the active site of a protein, and has been utilized to elucidate the virulence mechanism of a bacterial type III effector protein in its host cells (Fig. 3A).⁶⁸ A ruthenium-catalyzed cleavage reaction has been used for the cleavage of allyl carbamate to unmask caged Rhodamine 110 (R110) inside living cells (Fig. 3B).¹²⁸

Fig. 3 Transition metal-catalyzed decaging reactions under living conditions.

2.5. Selective labeling at cysteine residue

Cysteine is characterized by its remarkable nucleophilicity among 20 common amino acid residues. Traditionally, proteins can be site-specifically labeled at their solvent accessible cysteine residues by thiol-reactive alkylation reagents, e.g., maleimides and iodoacetamides.^69,129 However, excess maleimide-based reagents lead to the modification of the side chains of histidine, lysine and α-amino groups. By converting cysteine residues to dehydroalanine (Dha), the Dha residue can then be rapidly and specifically labeled by various thiol reagents.⁷¹ This approach has been used for the preparation of post-translationally modified proteins, including phosphorylation, glycosylation, methylation, acetylation, and lipidation (Table 1F).^130,131

Adjacent cysteines are usually oxidized to form disulfide bonds under non-reducing conditions. In these cases, the solvent accessible disulfide bond can be first gently reduced and subsequently “intercalated” by mono-sulfone reagents. This approach permits the site-specific PEGylation of a variety of therapeutic proteins, including human interferon α-2b and antibody fragments.⁷⁴ A more recent approach employed 1,3-dichloroacetone (DCA) to introduce a reactive ketone tag, enabling subsequent oxime ligation (Table 1F).⁷⁵

N-terminal cysteine displays unique reactivity. Proteins carrying an N-terminal cysteine can undergo native chemical ligation with thioester probes and chemoselective ligation with aldehydes to form thiazolidines.¹³² N-terminal cysteine can also specifically react with cyanobenzothiazole (CBT) derivatives at a fast reaction rate (9 M⁻¹ s⁻¹).⁷² The reaction of CBT compounds with D-cysteine is highly biocompatible and has been used for bioluminescent imaging of protease activity in live mice.¹³³

Palladium-tolyl complexes using 2-dicyclohexylphosphino-2′,6′-diisopropylbiphenyl (RuPhos) as the ligand have been developed to mediate efficient and highly selective cysteine conjugation reactions under biocompatible reaction conditions.⁷⁰ At pH 7.5, the rate of a palladium-medicated reaction is comparable to that of the maleimide reaction. This bioconjugation strategy has demonstrated its broad utility for making stapled peptides, site-specific labeling of proteins with a coumarin fluorophore, and the preparation of antibody-drug conjugates (ADCs) (Scheme 2).⁷⁰

Scheme 2 Organometallic palladium reagents for cysteine bioconjugation on proteins.

3. Chemoselective labeling of native proteins

Chemoselective labeling of native proteins is useful for the preparation of protein conjugates such as ADCs¹³⁴ and the labeling of endogenous proteins in living cells.¹³⁵ In addition to selective labeling at cysteine residues as discussed in the previous section, there are a few other approaches available (Scheme 3).

Scheme 3 Methods for chemoselective labeling of native proteins. LG: leaving group.

3.1. N-terminal labeling

The N-terminus of a protein shows unique reactivity and can be site-specifically labeled or converted to a bioorthogonal handle. The N-terminal α-amino group features a lower pK_a value (ca. 8) than that of the ε-amino group of lysine (ca. 10). Transamination methods have been used to convert the N-terminal α-amino group to an aldehyde or a ketone using glyoxylate in the presence of a divalent metal ion and a base¹³⁶ or using pyridoxal-5-phosphate (PLP).¹³⁷ N-terminal glycine shows the highest reactivity in the transamination reaction.¹³⁷ A ketene compound has been reported to selectively react with an N-terminal α-amino group at pH 6.3.¹³⁸ Another approach is the oxidation of an N-terminal serine or threonine to an aldehyde or ketone by sodium m-periodate at neutral pH.¹³⁹ Afterwards, the N-terminal aldehyde or ketone tag is ready for oxime ligation, Mukaiyama-Adol condensation, Pictet-Spengler reaction, etc., as discussed in previous sections (Scheme 3A).

3.2. Kinetically-controlled protein labeling (KPL)

Under normal bioconjugation conditions for protein labeling at lysine side chains, it is usually difficult to achieve site-specificity using regular amine-reactive reagents, such as an N-hydroxysuccinimidyl (NHS) ester. Nonetheless, lysine residues on a protein’s surface display subtle differences regarding their individual reactivity. This difference may originate from the different solvent accessibility of lysine, or the interaction of lysine with neighboring residues, or the combination of both.¹⁴⁰ Through kinetically-controlled protein labeling (KPL), proteins can be mono-modified at a specific lysine side chain.¹⁴⁰ This approach enables the site-selective introduction of a terminal alkyne, or azide into native proteins for a click reaction¹⁴⁰ or Staudinger ligation (Scheme 3B).¹⁴¹ KPL has been utilized for site-specific biotinylation of pharmaceutically relevant proteins for delivery into macrophages¹⁴² and dual labeling of the peptide hormone, somatostatin, for the visualization of targeted drug delivery in tumor cells.¹⁴³

3.3. Catalytic affinity labeling

Catalytic modules tethered with a ligand can be targeted to the binding pocket of a protein to locally catalyze the labeling reaction on the protein. For instance, a biotin-conjugated dimethylglycine (DMG) group is brought to the proximity of the avidin binding pocket, where it specifically catalyzes the acylation of amine probes with Asp108.¹⁴⁴ In the so-called modular affinity labeling (MoAL) approach, three modules are required: (1) the catalytic ligand module (biotin-DMG), (2) the labeling module (amine probe) and (3) the reactive module (CDMT). Another catalytic affinity labeling method is termed affinity-guided DMAP (4-dimethylaminopyridine) chemistry. DMAP is an effective acyl transfer catalyst, which can activate an acyl ester for its transfer to a nucleophilic residue. Therefore, a ligand tethered with DMAP allows specific labeling of native proteins by acyl ester probes (Scheme 3C).¹⁴⁵

3.4. Affinity labeling of endogenous proteins

In ligand-directed tosyl (LDT) chemistry, ligand-tethered tosyl ester probes are used to label endogenous proteins in cells. Upon labeling, the ligand is cleaved off at the tosyl ester linkage. Therefore, the protein is still active after labeling because the active center is no longer occupied by its ligand.¹⁴⁶ The concept was employed to construct a turn-on probe based on the release of the quencher upon labeling.¹⁴⁷ The LDT chemistry has been applied to the labeling of cell surface receptors¹⁴⁸ and native FKBP12 in live cells.¹⁴⁹ LDT chemistry is slow and typically requires over 12 hours of incubation time. Recently, a faster affinity labeling approach, known as ligand-directed acyl imidazole (LDAI) chemistry, was used to selectively modify the endogenous folate receptor at the cell surface (Scheme 3D). The experiments showed that LDAI labeling is 12-fold more efficient than LDT labeling.¹⁵⁰ New emerging affinity labeling reagents include chemical probes bearing an O-nitrobenzoxadiazole (O-NBD) unit. Upon the ligand-directed reaction with a lysine side chain, the non-fluorescent O-NBD is converted to fluorescent N-NBD. Translocator protein (TSPO) ligands carrying an O-NBD unit enabled proteomic identification of a partner protein of TSPO, a voltage-dependent anion channel (VDAC). The affinity labeling reaction using O-NBD is quite efficient with yields of 41% and 76% after 1 h and 12 h, respectively.¹⁵¹

4. Chemical labeling of proteins in living cells

Proteins function in signaling pathways and interacting networks in the complicated surroundings of cells and organisms. Fluorescent proteins (FPs) have revolutionized our ability to visualize and investigate protein function directly in living cells and organisms by fusion of individual proteins with FPs. Chemical probes, including organic dyes, are able to achieve properties that are not readily possible when using FPs. For instance, many organic dyes are superior to FPs in terms of brightness, photostability, far red-emission, environmental sensitivity, pulse-chase labeling and the flexibility for modifications to their spectral and biochemical properties. In the chemical tagging approach, a protein of interest (POI) is fused with a polypeptide tag, which is subsequently labeled with chemical probes. In general, these tags can be classified in the following categories: (1) metal chelation based peptide tags; (2) self-labeling peptide tags; (3) ligand binding domains (LBDs); (4) self-labeling enzymatic domains; (5) peptide sequences for enzymatic modifications; and (6) genetically encoded unnatural amino acids (UAAs) as “minimalist” tags (Table 4).

Table 4 Representative chemical tags for the labeling of proteins in cells

Tag name (size in AA)	Probe	Enzyme	Cellular labeling	Features	Reference(s)
AA: amino acids. (A) BG: O^6-benzylguanine; BC: O^2-benzylcytosine; BG-FL: BG-fluorescein conjugate; BC-FL: BC-fluorescein conjugate. (E) ACP: acyl carrier protein; PPTase: 4′-phosphopantetheinyl transferase; CoA: coenzyme A; PCP: peptide carrier proteins.
A. Metal chelation based peptide tags
His-tag (6): HHHHHH		No	Cell surface/intracellular	Ni(II) is toxic to cells and quenches fluorescence; intracellular labeling is possible using cell-penetrating multivalent chelator carrier complexes	Guignet (2004)^152
					Wieneke (2014)^153
His-tag (6): HHHHHH	Zn(II) complex: HisZiFit	No	Cell surface	Zn(II) is non-toxic; the zinc complex is membrane-impermeant	Hauser (2007)^154
D4 tag (4): (DDDD)n, n = 1–3	Multinuclear Zn(II) complexes: Zn(II)-DpaTyrs probe	No	Cell surface	Spectroscopic change upon chelation; D4 tag/Zn(II)-DpaTyrs pair is orthogonal to the His tag/Ni(II)-NTA pair	Ojida (2006)^155
B. Self-labeling peptide tags
CCXXCC (6) (tetracysteine)		No	Intracellular	Fluorogenic; non-specific reactions and interactions to endogenous thiols; biarsenical probes may be cytotoxic	Griffin (1998)^156
					Gaietta (2002)^157
SSPGSS (6) (tetraserine)		No	Cell surface	Fluorogenic; reduced background staining; cytotoxic; off-target labeling of endogenous tetraserine motif; RhoBo is cell permeable	Halo (2009)^158
HyRe tag (11): HKSNHSSKNRE		No	Cell surface	Neutral pH; imine byproduct formation at pH <6	Eldridge (2011)^159
SpyTag (13)	SpyCatcher protein domain (138 AA)	No	Cell surface	k = 1.4 × 10^3 M^−1 s^−1; high yield under diverse conditions of pH, temperature and buffer; formation of a stable amide bond linkage; not traceless	Zakeri (2012)^160
E3 tag (21): (EIAALEK)3	K3 peptide (21 AA): (KIAALKE)3	No	Cell surface	E3/K3 coiled-coil template induces proximity-driven rhodium(II)-catalyzed modification or acyl transfer reaction (t1/2 = 10 min)	Chen (2011)^161
					Reinhardt (2014)^162
C. Ligand binding domains (LBDs)
FKBP’ (108) (FKBP12_F36 V)		No	Intracellular	Specific binding between FKBP’ and SLF’; non-covalent	Clackson (1998)^163
					Marks (2004)^164
eDHFR (159)		No	Intracellular	Tight and specific binding between eDHFR and TMP; non-covalent/covalent; non-cytotoxic; affinity conjugation is rapid (t1/2 = 1.8 min)	Miller (2005)^165
					Liu (2014)^166
PYP-tag (125)		No; PYP tag is derived from a purple bacteria	Intracellular	PYP-tag covalently binds to the thioester derivative of cinnamic acid/coumarin; fluorogenic; up to k = 3950 M^−1 s^−1	Hori (2009/2013)^167,168
D. Self-labeling enzymatic domains
SNAP tag (182)		hAGT (mutated human O^6-alkylguanine-DNA alkyltransferase)	Intracellular	BG: k = 0.3 × 10^4 M^−1 s^−1; BG-FL: k = 2.8 × 10^4 M^−1 s^−1; removal of guanine after labeling	Keppler (2003)^169
CLIP tag (182)			Intracellular	BC-FL: k = 0.1 × 10^4 M^−1 s^−1. SNAP-tag and CLIP-tag possess orthogonal substrate specificities; removal of cytosine after labeling	Gautier (2008)^157
BL-tag (263)		Non-catalytic β-lactamase variant	Cell surface	k = 7.8 × 10^4 M^−1 s^−1, E166N mutant of β-lactamase; fluorogenic labeling	Mizukami (2009)^170
Halo tag (290)		Dha (mutated haloalkane dehalogenase)	Intracellular	k = 2.7 × 10^6 M^−1 s^−1; covalent; probe easily accessible by synthesis	Los (2008)^171
Cutinase (197)		A small globular serine esterase of 22 kDa	Cell surface	Cutinase is labeled by the covalent inhibitor, p-nitrophenyl phosphonate (pNPP)	Bonasio (2007)^172
E. Enzymatic modifications
ACP (77)/S6(12)/A1(12)/PCP(80)/ybbR tag(11)	Coenzyme A (CoA) derivatives	PPTase (AcpS or Sfp)	Cell surface	AcpS/Sfp catalyzes the transfer of a CoA-activated probe to the Ser residue of the tag. kcat = 5.5 × 10^−4–0.32 s^−1, kcat/Km = 5.3–4550 M^−1 s^−1	George (2004)^173
					Yin (2005)^174
					Zhou (2007)^175
AP tag (15): GLNDIFEAQK̲IEWHE		Biotin ligase (BirA)	Cell surface	Ketobiotin is a substrate for BirA and is incorporated into the Lys residue of the peptide tag; labeling time >20 min	Chen (2005)^83
Q-tag (7): PKPQQFM	Cadaverine derivatives	Transglutaminase (TGase)	Cell surface	TGase mediates the formation of an amide bond between the amine of cadaverine and the Glu residue of the Q-tag	Sato (1996)^176
					Lin (2006)^177
LAP tag (13–22)		Lipoic acid ligase (LplA)/LplA mutant	Cell surface/intracellular	k cat = 0.048 s^−1 for the conjugation of azido lipoic acid; coumarin addition by ^W37VLplA: kcat = 0.019 s^−1, kcat/Km = 3.8 × 10^2 M^−1 s^−1; ^W37ILplA: kcat = 0.016 s^−1, kcat/Km = 0.6 × 10^2 M^−1 s^−1	Fernandez-Suarez (2007)^178
LC̲XPXR (6)		Formylglycine generating enzyme (FGE)	Cell surface	FGE catalyzes the transformation of a Cys of the peptide tag to a formylglycine	Carrico (2007)^79
LPXT̲G (5)		Sortase A (SrtA)	Cell surface	SrtA cleaves the peptide at the Thr-Gly site, and attaches a labeled polyglycine peptide	Popp (2007)^179
CAAX (4)		Protein farnesyltransferase (PTFase)	Cell surface	CAAX is also present in endogenous proteins; an alkyne group is introduced for click labeling	Wollack (2009)^180
Split intein [Int^C (35) or Int^N (25)]	Another half of split intein [Int^N (100) or Int^C (104)]	Reconstituted split inteins	Cell surface/intracellular	Intein-mediated protein trans splicing removes intein after reaction (traceless); yield 55–90% and k = 0.9–4.7 s^−1 using Npu DnaE intein and AceL-TerL intein	Giriat (2003)^181
					Schütz (2014)^182
TITSS̲YYR (8)		AnkX from Legionella pneumophila	In vitro	k cat/Km = 0.9–5 × 10^2 M^−1 s^−1, covalent labeling can be removed by Lem1	Heller (2015)^183
Tub-tag (14): VDSVEGEGEEEGEE		Recombinant tubulin tyrosine ligase (TTL)	In fixed cell	Yield up to 99% with moderate enzyme concentrations and short reaction times	Schumacher (2015)^184
F. Incorporation of UAA as a “minimalist” tag
		BCN-RS/tRNACUA	Cell surface/intracellular	UAA bearing BCN or TCO can be site-specifically incorporated to proteins in live cells for labeling with tetrazine dyes	Lang (2012)^46

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4.1. Metal chelation based peptide tags

Metal chelation methods have been adopted for affinity chromatography in protein purification. The principle has also been utilized for protein labeling, either by non-covalent complex formation or by chelation-driven affinity conjugation.^152,185 Examples include the poly-histidine tag (His-tag, HHHHHH) and the tetra-aspartate tag [D4 tag, (DDDD)_n, n = 1–3], which can be labeled using Ni-NTA probes and zinc complexes, respectively.^{152,154,155,186–189} Specific intracellular labeling of His-tagged proteins was achieved using cell-penetrating multivalent N-nitrilotriacetic acid (NTA) carrier complexes.¹⁵³ The advantages of metal chelation based labeling can be attributed to the small size of the tag which confers minimal disturbance to protein function, the high labeling efficiency, selectivity and accessibility of various functional probes (Table 4A).¹⁹⁰

4.2. Self-labeling peptide tags

Tsien and coworkers reported the biarsenical FlAsH (fluorescein arsenical hairpin) as the first chemical surrogate to FPs for labeling proteins in live cells.¹⁵⁶ A red version of FlAsH, a resorufin-based biarsenical (ReAsH), was developed later. FlAsH and ReAsH probes allow specific and fluorogenic labeling of proteins fused to a tetracysteine motif (CCXXCC).^191–193 However, this labeling technology suffers practically from nonspecific labeling of thiol-rich biomolecules in the cell and the toxicity of the biarsenical ligands.¹⁹⁴ Nevertheless, the tetracysteine motif (CCXXCC) is much smaller than FPs. Other self-labeling peptide tags include the bisboronic RhoBo-tetraserine tagging system,¹⁵⁸ hydrazide-reactive (HyRe) tag,¹⁵⁹ SpyTag¹⁶⁰ and E3 tag^161,162 (Table 4B).

Prominent applications of self-labeling peptide tag approaches involve visualization of newly synthesized proteins and tracking of protein trafficking in live cells. This is readily achieved via the “pulse-chase” technique. The old populations of proteins were pulse-labeled by green-emitting FlAsH, while the newly synthesized proteins were chased by red-emitting ReAsH. Consequently, old and new copies of an individual protein were labeled using two colors. In one example, this approach was used to elucidate the mechanism of connexin assembly and turnover in HeLa cells.¹⁹¹ In another example, the approach was employed to study AMPA receptor (AMPAR) trafficking. Regulation of AMPA receptor (AMPAR) trafficking is important for neural plasticity. GluR1 and GluR2 are two AMPAR subunits that play a key role in the activity-dependent trafficking of the AMPARs during long-term potentiation (LTP) and depression (LDT). In order to examine the trafficking and synthesis of GluR1 and GluR2, a tetracysteine motif (EAAAREACCRECCARA) was attached at the C-termini. ReAsH-EDT₂ was first applied and after 6–8 h, FlAsH-EDT₂ was applied to cells expressing tetracysteine-tagged GluR1 or GluR2 (Scheme 4B). In this case, the red ReAsH-EDT₂ labels all preexisting GluR1/2 subunits, while the green FlAsH-EDT₂ labels those AMPRAR subunits synthesized during the 6–8 h chase period. The measurements suggested that both GluR1 and GluR2 are synthesized in dendrites and that an activity blockade enhances the dendritic synthesis of GluR1 but not GluR2.¹⁹⁵

Scheme 4 Dual-color “pulse-chase” labeling of neurons using ReAsH and FlAsH for the visualization of the synthesis and trafficking of AMPA receptors.

4.3. Ligand binding domains (LBDs)

The specific interaction between the ligand binding domain (LBD) and its small-molecule ligand confers specificity of labeling in cells. SLF’, a derivative of the synthetic ligand of FKBP12 (FK506 binding protein), binds to FKBP’ (FKBP12_F36 V mutant) with more than 1000-fold selectivity over the wild type FKBP12 protein.¹⁶³ Based on this specific binding, proteins fused with FKBP’ have been selectively labeled by SLF’-conjugated probes in live cells.¹⁶⁴ The antibiotic trimethoprim (TMP) is a specific inhibitor of E. coli dihydrofolate reductase (eDHFR). TMP binds to eDHFR with nanomolar affinity, which is over 1000-fold higher than the interaction with mammalian DHFR. As a result, the off-target labeling by TMP probes is minimal under certain conditions.¹⁶⁵

Because of the non-covalent binding, the labeling via the FKBP’ or eDHFR tag is reversible. In order to achieve a stable labeling, the affinity conjugation approach has been introduced. A cysteine mutation is introduced in the proximity of the TMP binding site on eDHFR, which can be specifically labeled by TMP-acryloyl probes due to a proximity-induced effect.¹⁹⁶ The reaction of a mildly reactive acryloyl group with other thiols in the cell is minimal under certain conditions.¹⁹⁷ Based on the affinity conjugation principle, a rapid and fluorogenic TMP-AcBOPDIPY probe is developed with a half-life of less than 2 min for covalent labeling (Fig. 4A).¹⁶⁶ Intracellular proteins fused with eDHFR_N23C were rapidly labeled by the TMP-AcBOPDIPY probe under no-wash conditions (Fig. 4B). In addition, the chemical probe displays a superior dynamic range in fluorescence lifetime imaging microscopy (FLIM) for intracellular FRET studies.

Fig. 4 (A) The chemical structure of the TMP-AcBOPDIPY probe (BOPDIPY: boron phenyldipyrrolemethene). (B) Cellular labeling of eDHFR_N23C fused with a K-Ras C-terminal sequence (CAAX), Rab1, Rab5, and nucleus localizing sequence (NLS) at the plasma membrane (PM), the Golgi body, the endosomes, and the nucleus, respectively, using the TMP-AcBOPDIPY probe under no-wash conditions. (C) The “tagging-then-labeling” approach using the TMP-AcAz ligand and BCN or DBCO probes for labeling intracellular proteins in live cells. (D) Specific labeling of EGFP-eDHFR_N23C/L28C-NLS at the nucleus in living HeLa cells by the BCN-TAMRA probe. Scale bar: 10 μm.

Recently, a more versatile “tagging-then-labeling” approach has been realized, enabling efficient introduction of bioorthogonal groups into proteins for bioorthogonal labeling in live cells. The TMP-AcAz ligand incorporates an azido group to proteins fused with the eDHFR tag (Fig. 4C). Subsequently, strain promoted cycloaddition reactions using DBCO- or BCN-conjugates facilitate protein labeling with various probes inside live cells (Fig. 4D).¹¹¹

The eDHFR tag has been used for live-cell imaging of protein–protein interactions (PPIs) between the first PDZ domain of ZO-1 (fused with eDHFR) and the C-terminal YV motif of claudin-1 (fused with GFP) using time resolved luminescence resonance energy transfer (LRET) technique (Fig. 5).¹⁹⁸ Conventional FRET imaging suffers from fluorescence breed-through, leading to high background. In the LRET approach, background signals from cellular auto-fluorescence and direct excitation of GFP were effectively eliminated by imposing a time delay of 10 μs between excitation and detection.

Fig. 5 Time resolved LRET for live-cell imaging of protein–protein interactions using the eDHFR tag.

Photoactive yellow protein (PYP) is a small (14 kDa) soluble protein found in several purple bacteria. It binds to a natural cofactor, the CoA thioester of 4-hydroxycinnamic acid through transthioesterification with Cys69. It also binds to the thioester derivative of coumarin-3-carboxylic acid. Since PYP and its ligands do not exist in animal cells, they can therefore be employed for bioorthogonal labeling of proteins (k = 1.1–124 M⁻¹ s⁻¹).^167,168

4.4. Self-labeling enzymatic domains

Enzyme-catalyzed reactions that proceed via irreversible conjugation with “suicide substrates” have been used for protein labeling in live cells. A variety of enzyme–substrate pairs are available. Many of these enzymes are able to tolerate modifications to their substrates. By generating fusion constructs with a protein, it is possible to covalently link the modified substrate to the protein. Generally, these reactions are specific, rapid and irreversible.

O ⁶-Alkylguanine transferase (AGT), a human DNA repair protein, has been used as a self-labeling tag.^169,199 The reaction involves the irreversible transfer of the alkyl group of O⁶-benzylguanine (BG) derivatives to the reactive cysteine residue within the enzyme to generate a covalently modified protein. More efficient AGT mutants, termed SNAP-tags (19 kDa), have been developed.¹⁶⁹ An orthogonal AGT-based tag, termed a CLIP-tag, reacts specifically with O²-benzylcytosine derivatives.¹⁵⁷

Single-molecule imaging often requires photo-stable and bright organic dyes, which are made possible using chemical protein labeling approaches. The spliceosome is a complex machine responsible for removing introns from the precursors of messenger RNAs (pre-mRNAs). The SNAP-tag has been exploited in combination with the eDHFR-tag to enable single-molecule imaging of the spliceosome in yeast cell extracts (Fig. 6A). The SNAP- and eDHFR-tags facilitate labeling pairs of the small nuclear ribonucleoprotein (snRNP) components of the spliceosome in cell extracts with bright organic dyes (TMP-Cy5 and BG-DY549), thereby enabling imaging of their assembly on individual pre-mRNAs. The measurements revealed that individual spliceosomal subcomplexes associate with pre-mRNA sequentially through an ordered pathway and that subcomplex binding is reversible.²⁰⁰

Fig. 6 (A) Experimental setup for single-molecule tracing of pre-mRNA splicing in yeast cell extracts using SNAP tagging and TMP tagging technologies. (B) Schematic depiction of HaloPROTAC in inducing degradation of HaloTag7 fused proteins. The chemical structure of one representative HaloPROTAC, HaloPROTAC-3 is given. Ub = ubiquitin; E3 = E3 ligase.

Halo-tag is a modified haloalkane dehalogenase that covalently binds to synthetic chloroalkane derivatives.^171,201 Halo-tag is commercially available and has been widely used to label proteins in cells. This approach has been applied to small molecule-induced protein degradation, namely, HaloPROTACs. PROTACs are a class of heterobifunctional molecules that link a ligand of E3 ligase to a ligand for a protein of interest (POI).²⁰² PROTACs recruit the E3 ligase to the POI, resulting in its ubiquitination and subsequent degradation by the proteasome. A bifunctional HaloPROTAC contains a chloroalkane and a hydroxyproline derivative which binds an E3 ligase VHL. The compound induces binding between the HaloTag7 fusion protein and the E3 ligase, leading to the degradation of the HaloTag7 fusion proteins by the proteasome (Fig. 6B).²⁰³

β-Lactamase is a small bacterial enzyme (29 kDa) that hydrolyzes β-lactam antibiotics. The E166N mutation of β-lactamase leads to the accumulation of acyl-enzyme intermediates due to the dramatic suppression of deacylation. The mutant β-lactamase, termed BL-tag, and β-lactam probes have been used for the covalent labeling of proteins in cells.¹⁷⁰ Enormous efforts that have been made on β-lactam antibiotics render it possible to design various β-lactam probes.²⁰⁴ Cephalosporin-based probes featuring substituent elimination facilitate the development of fluorogenic probes.²⁰⁵ Other examples in this category include cutinase¹⁷² and catalytic antibodies (Abs)²⁰⁶ (Table 4D).

4.5. Enzymatic modifications

A number of post-translational modifications have been harnessed to specifically incorporate chemical probes or bioorthogonal handles into proteins (Table 4E). The first example is the modification of cell surface proteins using E. coli biotin ligase (BirA). The enzyme recognizes and biotinylates a 15-mer acceptor peptide (AP) sequence. Using a synthetic ketone-containing biotin isostere (keto-biotin) as a substrate, BirA can be used as a “ketone ligase” to introduce a ketone handle for oxime ligation.⁸³ Another example is E. coli lipoic acid ligase, which is able to transfer a lipoic acid derivative carrying an azide¹⁷⁸ or a TCO moiety to proteins.¹¹⁹ The introduced azide and TCO moieties facilitate subsequent labeling using strained cyclooctyne and terazine probes, respectively.^119,178 The formylglycine-generating enzyme (FGE) specifically oxidizes the cysteine in the consensus LCXPXR motif to the formyl glycine. Subsequently, the aldehyde tag generated by the FGE on the target protein can selectively react with aminooxy and hydrazide probes.⁷⁹ Reconstitution of the split inteins mediates protein trans splicing (PTS). After ligation of the N- and C-peptide fragments, the intein is eventually removed.¹⁸¹ By choosing appropriate naturally split inteins, PTS reactions can proceed efficiently in live cells.^182,207 The bacterial enzyme AnkX from Legionella pneumophila has been used to transfer phosphocholine moieties from synthetically produced CDP-choline derivatives to a consensus sequence, TITSSYYR, at the termini or internal loop regions of a POI. The covalent labeling can be removed by another Legionella dephosphocholination enzyme Lem3.¹⁸³

Many other enzymes have also been used for the selective modification of proteins, including phosphopantetheine transferase (AcpS or Sfp),^{173–175,208} transglutaminases (TGases),^176,177 sortase (SrtA),¹⁷⁹ protein farnesyl transferase (PFTase),¹⁸⁰ glycosyltransferase,²⁰⁹N-myristoyl transferase (NMT)²¹⁰ and tubulin tyrosine ligase (TTL)¹⁸⁴ (Table 4E). The advantages of enzymatic modifications lie in the small size of the tag and the highly efficient reactions. However, many of the substrates are not cell permeable and therefore are not suited for intracellular labeling.

The enzymatic modification approach has been elegantly applied to spatially-resolved proteomic mapping in living cells.²¹¹ An ascorbate peroxidase (APEX) fused with a “mito” sequence was targeted to the mitochondrial matrix. Labeling was initiated by the addition of biotin–phenol and H₂O₂ to live cells. The resulting phenoxyl radicals are short-lived and membrane-impermeant and therefore only label neighboring endogenous proteins. The biotinylated proteins were recovered with streptavidin-coated beads and identified using mass spectrometry (Fig. 7). This approach led to the identification of 495 proteins within the human mitochondrial matrix, including 31 proteins that were not previously linked to mitochondria.

Fig. 7 Labeling the mitochondrial matrix proteome in living cells using mito-APEX and biotin-phenol.

4.6. Incorporation of UAA as a “minimalist” tag

Unnatural amino acid (UAA) mutagenesis has emerged as a powerful tool for the site-specific modification of proteins.²¹² The incorporation of UAAs into a protein sequence can be considered as the introduction of a “minimalist tag” (e.g. only an individual amino acid residue as opposed to a peptide sequence or a protein domain). Unnatural amino acids can be co-translationally incorporated into proteins in either a residue- or a site-specific fashion. The incorporation of UAAs carrying small bioorthogonal groups followed by chemoselective reactions makes it possible to label proteins with a diverse range of probes. A pool of UAAs featuring a variety of bioorthogonal handles has been added to the genetic codes of E. coli, yeast, and mammalian cells.³ This approach has been highly useful for super-resolution imaging, preparation of post-translationally modified proteins, and controlling protein dynamics and functions in living cells.^{46,68,117,213–215} This approach has also been used to make therapeutic proteins and to prepare a new generation of ADCs.¹³⁴ More details about this topic are summarized in a recent review.³

5. Concluding remarks

Selective protein labeling techniques have provided unprecedented views of protein structures, dynamics and functions in vitro, in live cells, and in whole organisms. These approaches have demonstrated the power of chemistry as useful tools in a wide range of biological research areas, including post-translational modifications of proteins, preparation of protein-based pharmaceuticals, super-resolution imaging,^117,216 visualization of intracellular protein–protein interactions, modulation of protein function in live cells, proteome labeling⁴⁹ and affinity-based protein profiling.¹¹⁴

Despite successful applications in selective protein labeling, numerous challenges remain in bioorthogonal chemistry. Firstly, many bioorthogonal functional groups are not truly “bioorthogonal”. For instance, strained alkynes may react with free thiols in live systems. Aldehyde and ketone functionalities are also present in many metabolites in living systems. Secondly, some of the bioorthogonal groups are too large and lipophilic, such as cyclooctynes and cyclooctenes, causing non-specific staining and reduction of effective reactants in live cells. Thirdly, the instability of tetrazine, phosphine and other groups in live cells during a prolonged incubation time leads to the deactivation of the bioorthogonal moieties. Fourthly, some metal-catalyzed reactions are incompatible with living conditions, as exemplified by the cytotoxicity of the copper ion used for CuAAC. To address these issues, the development of new bioorthogonal chemistry is definitely required. Since there are few “perfect” bioorthogonal reactions available, one has to carefully consider both the pros and cons of each reaction in order to identify the most suited ones for a particular application.

Chemical tagging approaches allow for the incorporation of probes or bioorthogonal handles into proteins in cells and organisms. However, most of these approaches rely on exogenous expression of the POI with tags or UAAs. Although the investigation of exogenous proteins is useful for unraveling biological processes, the advance in selective labeling of endogenous proteins should facilitate the proteomic analysis of cellular organelles and protein complexes, target identification and diagnosis. An advantage of chemical probes over FPs lies in the flexibility of modifications on organic dyes. Therefore, the development of new organic dyes with special properties, e.g. photo-switchable, activatable, highly fluorogenic, bright, far-red emissive, etc., should substantially help to understand the biological mechanisms of proteins in the context of living systems.

Acknowledgements

This work was supported by DFG grants (grant no. SPP 1623 and SFB 642 to Y. W. W.).

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