Labeling and Single-Molecule Methods To Monitor G Protein-Coupled


Jun 24, 2016 - In single-molecule fluorescence experiments, the primary role of the fluorophore is, of course, to reveal the existence of the molecule...
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Labeling and Single-Molecule Methods To Monitor G ProteinCoupled Receptor Dynamics He Tian,† Alexandre Fürstenberg,† and Thomas Huber* Laboratory of Chemical Biology and Signal Transduction, The Rockefeller University, 1230 York Avenue, New York, New York 10065, United States ABSTRACT: The superfamily of G protein-coupled receptors (GPCRs) mediates a wide range of physiological responses and serves as an important category of drug targets. Earlier biochemical and biophysical studies have shown that GPCRs exist temporally in an ensemble of interchanging conformations. Single-molecule techniques are ideally suited to understand the dynamic signaling and conformational complexity of G protein-coupled receptors (GPCRs). Here, we review the progress in single-molecule studies on GPCRs. We introduce the fundamental technical aspects of single-molecule fluorescence. We also survey the methodologies for labeling GPCRs with biophysical probes, particularly fluorescent dyes, and highlight the relevant chemical biology innovations that can be instrumental for studying GPCRs. Finally, we illustrate how the optical techniques and the labeling schemes have been combined to investigate GPCR signaling and dynamics at the single-molecule level.

CONTENTS 1. Introduction 2. The Era of Single Molecules in Biology 2.1. Why Single Molecules 2.2. Basic Requirements 2.3. Fluorescence Observables 2.4. Microscope Configurations: Spectroscopy or Imaging? 2.5. Observing Diffusing Molecules 2.6. Imaging Immobilized Molecules 2.7. Single-Molecule Trapping 2.8. Super-Resolution Imaging 2.9. Multicolor Single-Molecule Detection 3. Cell Biology and Biochemistry of GPCRs 3.1. GPCRs as Important Drug Targets 3.2. Assembly of the GPCR Signaling Complex 3.3. Spectroscopic and Structural Studies on GPCR Activation 3.4. Conformational Diversity of GPCRs 3.5. Membrane Dynamics and Oligomerization of GPCRs 4. Labeling of GPCRs with Biophysical Probes 4.1. Overview 4.2. How to Specifically Target a GPCR 4.3. Immunofluorescence Using Antibodies and Nanobodies 4.3.1. Antibodies 4.3.2. Nanobodies 4.4. Fluorescent Ligands 4.5. Ligand-Directed Labeling 4.6. Fluorescent Proteins and Fö rster Resonance Energy Transfer (FRET)

© XXXX American Chemical Society

4.7. Luciferase and Bioluminescence Resonance Energy Transfer (BRET) 4.8. Peptide-Based Tags 4.8.1. Arsenical Hairpin Binders Specific for the Tetracysteine Tag 4.8.2. Bisboronic Probe Specific for the Tetraserine Tag 4.8.3. Tetranuclear Zinc(II) Probe Specific for the Oligo-aspartate Tag 4.8.4. Template-Directed Labeling Based on a Coiled-Coil Motif 4.9. Chemoenzymatic Labeling Based on SelfLabeling Protein Tags 4.9.1. SNAP-Tag and CLIP-Tag 4.9.2. Halo-Tag 4.9.3. TMP-Tag 4.10. Chemoenzymatic Labeling Based on Posttranslational Modification Enzymes 4.10.1. Biotin Ligase and Lipoic Acid Ligase 4.10.2. Sortase 4.10.3. Formylglycine-Generating Enzyme 4.10.4. Ascorbate Peroxidase 4.10.5. Applications of the Engineered Posttranslational Enzymes 4.11. Classic Approach for Site-Specific Labeling of GPCRs 4.11.1. Targeting the Naturally Occurring Functionalities in GPCRs

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Chemical Reviews 4.11.2. Spectroscopic Studies on GPCRs Enabled by Cysteine and Lysine Labeling 4.11.3. Limitations to Targeting Naturally Occurring Functionalities 4.12. Novel Approaches for Site-Specific Labeling of GPCRs 4.12.1. Incorporating Unnatural Amino Acids into GPCRs 4.12.2. Genetically Encoded Unnatural Amino Acids as Biophysical Probes for GPCRs 4.12.3. Bioorthogonal Labeling of GPCRs Targeting Genetically Encoded Reactive Handles 4.12.4. Potential Issues with Amber Codon Suppression in Living Cells 4.13. Fluorogenic Labeling Reactions 4.14. Choosing the Right Labeling Method To Understand the Biochemistry and Cell Biology of GPCRs 4.14.1. Tracking GPCR Conformational Change 4.14.2. Trafficking and Internalization 4.14.3. Oligomerization 5. Application of Single-Molecule Methods to GPCRs 5.1. Mobility, Oligomerization, and Stoichiometry 5.2. Membrane Organization beyond the Diffraction Limit 5.3. Conformational Dynamics 5.4. Ligand Binding 5.5. Structure and Stability 6. Conclusion and Prospect Author Information Corresponding Author Author Contributions Notes Biographies Acknowledgments References

Review

facilitate the endeavor of drug discovery since a large proportion of existing therapeutic agents target GPCRs.1,2 Given the challenge of identifying effective drug targets, this classic approach to modulating GPCRs is unlikely to be ever out of fashion. To quote James Black, the British pharmacologist and Nobel laureate who was best known for developing βblockers targeting β-adrenergic receptors, “the most fruitful basis of the discovery of a new drug is to start with an old drug.” In the past decade, the GPCR field has witnessed an explosion of crystal structures, which has tremendously advanced the understanding of the structure−function relationship of the receptors.3,5 Crystallography is the reference method for providing high-resolution spatial information. Nonetheless, due to their static nature, crystal structures can only offer very limited insight into the dynamics of conformations and of signaling. Intrinsically, the crystallization of a protein eliminates its conformational diversity. Moreover, the extensive protein engineering required for crystallization often sacrifices the structural information on the flexible loops and termini. All of these considerations point to an acute need for developing alternative methodologies. Single-molecule techniques nicely complement crystallography in uncovering the dynamics and conformational complexity of the GPCR signaling complex. At the beginning of this Review, we describe the basic concepts in single-molecule fluorescence techniques. We also introduce several key events in GPCR signaling including the assembly of the GPCR signaling complex, receptor activation, receptor conformational diversity, and receptor oligomerization. A major technical challenge for single-molecule fluorescence experiments is to prepare suitable fluorescently labeled receptors. We therefore strive to provide an overview of the methodologies for attaching extrinsic probes to GPCRs, with an emphasis on fluorescent labeling. The enabling role of labeling strategies in spectroscopic and imaging experiments is illustrated, and the advantages and limitations of each strategy are discussed and compared. Many labeling schemes have been employed in ensemble measurements and may serve as the stepping-stones for single-molecule studies. We also examine recent innovations from chemical biology, such as liganddirected labeling, chemoenzymatic labeling, unnatural amino mutagenesis, bioorthogonal chemistries, fluorogenic reactions, and how these approaches can be transferred to the GPCR field. Relevant reviews are referenced to orient the GPCR specialists to explore the chemical biology toolkit. Finally, we review existing single-molecule studies on GPCRs and list them in an extensive table, with annotations specifying the techniques and summarizing the key findings.

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1. INTRODUCTION The G protein-coupled receptors (GPCRs) constitute a large family of transmembrane receptors that transduce extracellular signals into intracellular biochemical responses. GPCRs mediate a myriad of fundamental physiological processes, such as vision, smell, taste, neurotransmission, immune response, mood and behavioral regulation, homeostasis, metabolism, and energy balance. Three Nobel prizes have been awarded to scientists working on GPCRs: George Wald, who elucidated the biochemistry of the visual photoreceptor rhodopsin (physiology or medicine, 1967); Richard Axel and Linda Buck, who discovered olfactory receptors (physiology or medicine, 2004); and a few years ago Robert Lefkowitz and Brian Kobilka, who made groundbreaking contributions to understanding the molecular basis of GPCR function, in particular, that of adrenergic receptors (chemistry, 2012). Several other Nobel prizes were closely related to the GPCR signaling pathway. Understanding with molecular precision how GPCRs function in cells is an active area of research. In addition to a basic understanding of transmembrane signaling, studies of GPCRs can provide insights that might

2. THE ERA OF SINGLE MOLECULES IN BIOLOGY Single-molecule observation and manipulation in vitro and in living cells have been revolutionizing our way to address biological questions, enabling scientists to monitor elementary biochemical reactions with an unmatched level of detail and to capture images of ongoing processes in living cells at an unprecedented resolution. More than 25 years ago, the first optical detection experiments on single molecules, first at 4 K,6 then at room temperature,7 were mostly driven by the curiosity of physicists. Since then, single-molecule spectroscopy and imaging have emerged as routine techniques in biological research.8 Many areas have benefited from the information-rich high-quality data, which can nowadays be collected both on B

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readers of some basic requirements (Figure 2). Single-molecule fluorescence measurements traditionally require a transparent,

purified components in vitro or by following individual molecules in living cells.9 2.1. Why Single Molecules

Populations of biomolecules are most of the time heterogeneous. For example, the individual copies of the same protein or nucleic acid strand are in different conformations or at different stages of the enzymatic cycle. The main motivation for using single-molecule methods is to measure the full distributions of behavior and to expose hidden heterogeneities that would be totally obfuscated if only the population average were evaluated (which is the case in any measurement of an ensemble of molecules). The shape of the distribution might be skewed or reveal distinct subpopulations of molecules (Figure 1), potentially offering mechanistic insight. Furthermore, singleFigure 2. Excitation of a single molecule. (a) Typical Jablonski diagram for single-molecule fluorescence spectroscopy and imaging. A typical organic fluorescent probe is excited into vibrationally excited levels of its first excited state. After fast vibrational relaxation (VR) to the ground vibrational level of the excited electronic state, the molecule may either emit fluorescence, relax nonradiatively to its ground state via internal conversion (IC), or undergo intersystem crossing (ISC) to a nonemissive triplet state. (b) Schematic illustration of a focused optical beam exciting a single molecule on a coverslip surface. Key parameters for a good single-molecule fluorescent probe are molar extinction coefficient (ε), fluorescence quantum yield (Φfl), photostability, characterized by its photobleaching quantum yield (Φb), blinking characteristics, and linkability, that is, the ability to be selectively attached to molecules of interest.

Figure 1. Populations in single-molecule spectroscopy. Fluorescent lipids freely diffuse in a bilayer (left) but sometimes stop at a particular spot before starting diffusing again. An analysis of their motion reveals two populations: one with a high diffusion coefficient corresponding to the moving molecules, and another with a low diffusion coefficient corresponding to the immobilized molecules. A simple measurement of the average diffusion coefficient would, however, yield a value falling somewhere between the two populations, failing to represent the true molecular behaviors. Scale bar in the left panel: 5 μm.

nonfluorescent host matrix (crystal, polymer, solvent, cell) deposited on a thin coverslip made out of low fluorescence glass. In such a matrix, the molecules are observed at concentrations low enough for individual molecules to be separated in space (more than the diffraction limit of ∼200 nm), time, or wavelength.23 Apart from a few exceptions, biological molecules are optically undetectable unless coupled to a probe. A useful probe, most of the time a fluorophore, should have the following general properties: (1) absorbing light efficiently (high extinction coefficient); (2) emitting light efficiently (high fluorescence quantum yield); (3) displaying high photostability (the total number of photons emitted before photobleaching is large); and (4) specifically linkable to the molecule of interest. A steady flow of emitted photons with rare “blinking” events (fluorescence intermittence) is also highly desirable for most applications, except for singlemolecule based super-resolution imaging. Selective excitation is usually achieved using a laser beam resonant with the optical transition of the probe. Cellular autofluorescence limits singlemolecule detection to excitation with wavelengths typically larger than about 480 nm.

molecule measurements can follow the internal states of the same molecule over time and the transitions between them, thus revealing rare intermediate states or hidden kinetic pathways and circumventing the need for sample synchronization. Stochastic multistep processes, such as the dance of individual molecular motors or tRNA transit on single translating ribosomes, could be monitored.10,11 Typical singlemolecule labels behave like nanometer-sized light sources that are strongly influenced by their close surroundings. They are therefore local reporters of the local environment and provide a direct window into the nanoscale and its changes over time. A surprising observation from the early single-molecule experiments was precisely the changes in the fluorescence excitation or emission spectrum of a single molecule over time, a phenomenon termed “spectral diffusion” indicative of variations in the proximal environment of the probe.12−14 Remarkably, optical excitation enables observation of exactly one molecule surrounded by innumerable other transparent molecules composing a crystal, a polymer matrix, the solvent, or a cell. Therefore, single-molecule spectroscopy reaches the ultimate limit of sensitivity of ∼1.66 × 10−24 moles of the molecule of interest (1.66 yoctomoles). Being able to discriminate the signal arising from one, two, or a few molecules, single-molecule spectroscopy and imaging have found quantitative applications, including molecular counting or the determination of molecular stoichiometries.

2.3. Fluorescence Observables

In single-molecule fluorescence experiments, the primary role of the fluorophore is, of course, to reveal the existence of the molecule of interest as well as its position (Figure 3a). Multiple fluorophores of different colors can be simultaneously detected by sorting the emitted photons into different channels on the basis of their wavelengths. This multicolor detection scheme can be used to monitor colocalization of different molecules (Figure 3b). The location of a single fluorophore can be determined with nanometer precision in the laboratory frame (Figure 3c), enabling its trajectories to be reconstructed and the diffusional property to be evaluated.

2.2. Basic Requirements

The fundamental principles of single-molecule optical spectroscopy and imaging have been described extensively in many excellent reviews.8,9,15−22 Here, we will briefly remind the C

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Figure 3. Different processes that can be monitored by fluorescence spectroscopy and imaging at the single-molecule level. (a) A fluorophore (green) attached to a macromolecule reports its location, which can be determined within a few tens of nanometers. (b) Two molecules labeled with two different nonoverlapping fluorophores can be colocalized (green and red). (c) Localizing a single molecule over time can be used to track its moving trajectory and evaluate its diffusion properties. (d) The fluorescence intensity (I) of a single molecule going in and out of the excitation/ detection volume fluctuates over time (t) between a defined fluorescence level (arbitrary unit 1) and the background level. (e) Single-step photobleaching of noninteracting fluorophores enables the stoichiometry to be determined by counting the steps. (f) The orientational motion of a tethered fluorophore can be determined by using polarized excitation and detecting the polarization of the emission in two orthogonal channels (s and p). (g) Changes in the local environment of a fluorophore lead to fluctuations in the fluorescence intensity as well as lifetime. (h) Photoinduced electron transfer (PET) between a fluorophore and a quencher (Q) also modulates the fluorescence signal. (i) FRET between a donor (D) and an acceptor (A) fluorophore leads to anticorrelated changes in the donor and acceptor emission intensities.

Observing the intensity of a fluorescent spot over time can be informative by itself. For example, the binding dynamics of a fluorescent molecule to its immobilized partner can be monitored (Figure 3d). Similarly, the stoichiometry and the copy number of molecules within a complex can be determined by following stepwise photobleaching events and comparing the total intensity of a complex with the intensity of a single fluorophore (Figure 3e).24,25 In addition to the fluorescence intensity, other fluorescence parameters can provide valuable information on the dynamics of biomolecules.26 First, the polarization of the emission of single fluorophores reflects their rotational freedom (Figure 3f) that can be altered by intermolecular interaction or changes in its local environment (e.g., viscosity). Second, the fluorescence lifetime (typically a few nanoseconds) of a single molecule also changes with the local environment of the dye. It is worth noting that a change in fluorescence lifetime is commonly, but not necessarily, correlated with changes in the fluorescence intensity.14,27 Because more than one mechanism may change in the fluorescence intensity and lifetime, the interpretation is

not always straightforward (Figure 3g). An important mechanism is photoinduced electron transfer (PET), through which the lifetime of a fluorescent molecule can be shortened by a proximal quencher moiety (Figure 3h).28−30 Some amino acid residues, such as tyrosine or tryptophan, can serve as such PET quenchers. Therefore, fluorescence lifetime is useful in probing protein intramolecular conformational changes or intermolecular interaction.31 Finally, if two different fluorophores come close enough, dipole−dipole interaction between them results in Förster resonance energy transfer (FRET) (Figure 3i). Because the efficiency of the energy transfer process is sensitive to the distance between the donor and acceptor, single-molecule FRET (smFRET) is a powerful tool for detecting the conformational changes or molecular interactions involving macromolecules. We will further discuss smFRET in section 2.9. 2.4. Microscope Configurations: Spectroscopy or Imaging?

Single-molecule experiments are generally performed on inverted fluorescence microscopes configured either in confocal D

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Figure 4. Typical microscope configurations in single-molecule fluorescence detection experiments.

Figure 5. Fluorescence correlation spectroscopy (FCS) and single-particle tracking (SPT). (a) FCS of single diffusing molecules. Diffusion of fluorescent molecules through an excitation laser beam leads to fluctuations in the fluorescence intensity over time. An autocorrelation analysis of the fluorescence intensity time trace reports on the time scales of the fluctuations and enables one to extract the diffusion coefficient τD of the studied species. (b) SPT. Movies of diffusing single molecules are recorded and the position of each molecule is subsequently determined with subpixel resolution in every frame. Determining positions of single molecules throughout the frames enables one to analyze individual trajectories and to extract the mean squared displacement (MSD) of the individual molecules. The shape of the MSD curve depends on the diffusing behavior of the molecule of interest and can, for example, distinguish cases where a molecule undergoes pure random Brownian motion (red) or is confined to a given area (green).

charge coupled device (EMCCD) or a scientific complementary metal−oxide−semiconductor (sCMOS) camera, to directly generate an image without the need to scan. Many fluorophores can be observed with low excitation intensity (typically on the order of a few W/cm2) at video rate, enabling translation of single molecules to be observed in real time. In a standard epifluorescence configuration, a large volume of the sample is excited so that background signal from out-of-focus emitters may severely affect imaging quality, especially for thicker samples. This problem is solved by the total-internal-reflection fluorescence (TIRF) configuration, in which a region less than 0.1 μm in thickness (as compared to 100 μm, or more, in epifluorescence) is selectively excited by the evanescent field of a totally internally reflected laser beam entering the objective off-center. TIRF is, however, only useful when the molecules of interest are immobilized on the surface of the coverslip, or confined to a narrow space above it. The confocal configuration achieves a greatly enhanced signal-to-noise ratio relative to wide-field imaging. The diffraction-limited illumination volume results in high excitation intensity (typically a few kW/cm2 on average), while the introduction of a pinhole greatly reduces the background. Furthermore, the temporal resolution of PMT or APD detectors is orders of magnitude better than that of cameras used in wide-field illumination, whose temporal resolution is limited to milliseconds by the necessity to read out a twodimensional detector array. Therefore, confocal point detection is preferred in spectroscopy experiments on single molecules. However, when it comes to observing dynamic structures, imaging large regions of interest, or tracking single molecules over time, wide-field configurations are clearly advantageous

scanning or in wide-field illumination mode (Figure 4). An infinity-corrected, high-numerical-aperture objective lens is used to excite the sample as well as to collect fluorescence emission. High-quality optical beam splitters and filters are required to separate the excitation and emission light, because the emitted photon flux from single fluorescing molecules is many orders of magnitude weaker than that of the excitation light.17 In a confocal configuration, the sample is illuminated with the smallest possible spot, which is achieved by focusing a collimated laser beam with the microscope objective to produce a diffraction-limited spot at the sample plane. The size of the illumination spot depends on the wavelength (λ) of the light and on the numerical aperture (NA) of the objective (a description of how much light the objective is designed to collect; typical oil-immersion objectives have NA values between 1.4 and 1.5). The dimension of the illumination spot can be estimated using Abbe’s limit for spatial resolution (∼0.5λ/NA, approximately 200 nm for visible light). The collected filtered emission is focused through a pinhole, which rejects out-of-focus light and thereby minimizes background photons that reach a point detector, like an avalanche photodiode (APD) or a photomultiplier tube (PMT). Moving the excitation light or the sample can cover a larger region of interest. Another class of single-molecule techniques uses wide-field microscopy. Here, the excitation source (usually a laser) is focused onto the back aperture of the objective to illuminate an area typically tens of micrometers in diameter. Fluorescence photons across the illuminated region are collected with a twodimensional array detector, such as an electron-multiplying E

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recognizes epitope-tagged molecules of interest, for example, GPCRs (Figure 6).39−41

because they allow different regions of the sample to be imaged simultaneously in real time. 2.5. Observing Diffusing Molecules

Molecules are not static in solution; they undergo constant Brownian motion. When a fluorescent molecule diffuses through a focused (confocal) laser beam, it gives a short fluorescence burst that lasts only up to a few milliseconds. By calculating the autocorrelation of the fluorescence intensity fluctuations over time arising from diffusing molecules, fluorescence correlation spectroscopy (FCS, Figure 5a) extracts information about average diffusion coefficients and molecular rotation predominantly at the origin of these fluctuations. Because the intensity of the autocorrelation function depends on the number of molecules present in average in the confocal volume, typically about one femtoliter, concentrations of samples in the picomolar to the nanomolar range can be determined by FCS.32 Although at a sufficiently low concentration the fluorescence fluctuations recorded in FCS experiments truly arise from single molecules, the outcome of such measurements is still the average value describing the entire ensemble. To characterize the underlying distribution of diffusional behavior, single fluorescent molecules are followed in real-time by wide-field fluorescence microscopy using the single-particle tracking (SPT) technique (Figure 5b).33 The fluorescent molecules are localized in a series of recorded frames. The two- or threedimensional trajectories can be reconstructed by comparing the adjacent frames. Diffusion coefficients are extracted by evaluating the mean squared displacement (MSD) of the particle over time. Such diffusion analysis can be used to classify the type of motion that the fluorescent molecule has undergone: purely Brownian (freely diffusing), directed (as is the case when there is flow or if a protein travels along the cytoskeleton of a cell), confined to an area, or immobile. The comparison of many individual trajectories of one type of molecules has given insights into the spatial or temporal heterogeneities of biological processes. For example, the diffusion coefficient of glycine receptors was found to depend on their location in the neuronal somatodendritic membrane and to change over time.34

Figure 6. Example of a general surface passivation and capturing scheme. The coverslip surface is coated with a silane layer to which biotinylated BSA is then cross-linked. The biotin (B) can bind an avidin protein (A), which in turn recognizes a biotinylated antibody. The latter recognizes a specific epitope of the protein of interest, in the case of this cartoon a GPCR embedded in a detergent micelle and sitespecifically labeled with a single fluorophore.

A second widely used passivation method involves silanization of the glass surface. Typical bifunctional silanization reagents bind covalently to the glass through an organofunctional alkoxysilane group. A large variety of organofunctional groups are available for modifying the surface property of glass. For example, aliphatic hydrocarbon or polyethylene glycol/ polyoxyethylene (PEG/POE) chains render the surface hydrophobic or hydrophilic, respectively.42,43 POE detergents, such as Tween-20, are used together with hydrophobic silanes to prepare hydrophilic glass surfaces,44 which efficiently prevent sticking of some biomolecules. These hydrocarbon or PEG/ POE chains can be further functionalized with chemically reactive groups to attach biomolecules of interest by amine- or thiol-selective reactions.

2.6. Imaging Immobilized Molecules

The time scale of biological reactions varies from milliseconds to seconds, and it is often desirable to observe a stationary molecule as long as possible. Several strategies have been developed to immobilize molecules to the surface of a coverslip.35 One way is to embed them into a polymer matrix such as gelatin, poly(vinyl alcohol) (PVA), or poly(methyl methacrylate) (PMMA).36,37 However, many biomolecules exhibit natural affinities for glass and can be captured on a surface simply by immersing a precleaned coverslip in a dilute solution. The problem is that impurities may also stick to the coverslip surface. Different methods have been devised to prevent nonspecific binding. Typically, a bifunctional passivation layer is required, which on one hand blocks the glass surface and prevents undesirable binding of nontarget molecules, and on the other hand specifically captures the target molecules. A popular choice of passivation layer is biotinylated bovine serum albumin (BSA) that nonspecifically adsorbs to the glass.38 Biotin binds to tetravalent avidins with high affinity and very slow dissociation rate. Avidins serve as the intermediate layer to connect the biotinylated BSA and the biotinylated target, or a biotinylated antibody that specifically

2.7. Single-Molecule Trapping

Surface immobilization by chemical means enables prolonged observation of biomolecules, but doubts persist regarding whether surface-attached molecules behave the same as molecules freely diffusing in solution. When the aim is to monitor subtle conformational changes and dynamics, additional control experiments are often required. The antiBrownian electrokinetic (ABEL) trap developed by Cohen and Moerner has made it possible to hold individual biomolecules in solution for up to several seconds without altering their internal degrees of freedom (Figure 7).45−47 It combines fluorescence-based position estimation with fast electrokinetic feedback to counter Brownian motion of a single F

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reduces the size of the microscope point spread function (PSF) by using patterned excitation beams and nonlinear response effects.56,57 The second class of imaging methods based on localization of single photoswitchable fluorescent molecules has become particularly popular due to their relative ease of implementation and the nature of qualitative and quantitative answers they can offer.58−61 These methods may be regrouped under the term of single-molecule localization microscopy (SMLM, Figure 8).62

Figure 7. Principle of the ABEL trap. In conventional FCS experiments, molecules freely diffuse in solution and give rise to very short fluorescence bursts as they pass through the excitation beam (trap off). When the electrokinetic feedback of the ABEL trap is turned on, individual fluorescent molecules are maintained within the excitation volume in a microfluidic cell (left) due to fast position readout and application of voltage pulses via four electrodes. Figure adapted and reproduced from ref 48. Copyright 2012 American Chemical Society.

molecule. The molecule is thereby maintaining its position in the field of view. ABEL trapping can be viewed as a form of real-time electrophoresis. Such a trap also applies to noncharged molecules of interest by taking advantage of the electroosmotic effect, that is, the ability of the trap to create a flow by dragging ions in solution.48 Because the molecule is kept in a region of time-averaged uniform intensity, the fluctuations in the fluorescence intensity can be informative. Conformational dynamics of DNA and proteins on the millisecond to second time scale, as well as molecular stoichiometries, have been successfully monitored with an ABEL trap.27,49,50 Interestingly, the photodynamics of allophycocyanin protein molecules trapped in solution differed substantially from the cases where the protein was either attached to a surface or embedded in polymers.51

Figure 8. Principle of single-molecule localization microscopy. (a) Fluorophores must be photoswitched between a dark and an emitting state. (b) Single molecules can be localized in individual frames and their position determined with nanometer accuracy by approximating their intensity profile by a mathematical function (usually a twodimensional Gaussian). The scale bar represents 1 μm. (c) By repeating the process over many thousands of frames, a superresolution image can be reconstructed from the list of molecular coordinates in a pointillist fashion.

The principle underlying SMLM is the realization that the center of the PSF originating from a single molecule (the “peak” of the fluorescence “mountain”) can be localized with much better accuracy than the width of the PSF itself.63,64 The localization precision (σ) of a single emitter is statistically related to the number of photons (N) detected from that emitter (σ ∝ N−1/2). Typically, the localization precision is on the order of 10−50 nm. SMLM relies on three basic principles to obtain a super-resolved image (Figure 8): first, fluorescent probes that can be photoswitched actively or stochastically between a fluorescent and a dark state; second, the repeated detection of many isolated emitters; and third, the reconstruction of a super-resolution image from the singlemolecule localization data. In each imaging frame, only a low number of molecules is excited to the fluorescent state so that single emitters can be spatially resolved and localized despite a high density of labeling. These simple principles can be implemented in various ways,62,65,66 and have been used to obtain static and dynamic structures of many biological and nonbiological samples.67,68 SMLM can also be used to determine the relative or absolute molecular copy numbers and cluster sizes.69,70 Furthermore, the direct output of SMLM is not an image but a list of molecular localizations and intensities. This form of data intensities has enabled direct coordinate-based data analysis schemes that would be impossible to realize with the pixelated intensity information generated by conventional optical microscopy.69,70

2.8. Super-Resolution Imaging

The size of a typical fluorescent label may vary from ∼1 nm for a single organic fluorophore, to 2−3 nm for a fluorescent protein, and up to 6−8 nm for the core of a luminescent nanocrystal (“quantum dot”). No matter how good the optical microscope is, all of these point-like objects appear at least ∼200 nm in size. This spot size in an image is determined by the diffraction of light that fundamentally limits the resolution (d) to about one-half the wavelength (λ) of the light (see eq 1), even with high-numerical-aperture (NA = n sin θ) optics. Initial attempts to achieve subdiffraction resolution involved near-field imaging, which used an aperture much smaller than the wavelength of light and thus only detects the emission light leaking through this tiny hole. Near-field scanning optical microscopy (NSOM) yielded the first success in imaging a single fluorescent molecule at room temperature.52 Nonetheless, the widespread application of NSOM has been hampered by its fundamental limitations related to the low-intensity light throughput through the very small aperture, the difficulty of implementation, and the actual short-range of the near-field that prevented it from becoming a widespread subdiffraction imaging technique.53 d = λ /(2n sin θ )

(1)

Two other approaches for overcoming the diffraction limit of light were theorized in the mid-1990s54,55 and experimentally demonstrated about a decade later, resulting in the recent boom of so-called super-resolution methods. The first class, exemplified by stimulated emission depletion (STED) microscopy and structured illumination microscopy (SIM), directly

2.9. Multicolor Single-Molecule Detection

An early development crucial for addressing increasingly complex mechanistic questions was the introduction of multiple G

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excitation wavelengths and multiple detection channels. Multichannel detection can nowadays easily be achieved using appropriate lasers and filter sets. Multicolor imaging has been applied to all methods described above. In particular, highresolution colocalization of single molecules can be achieved by imaging two different types of fluorophores separately and localizing them to nanometer accuracy.71,72 Both intermolecular and intramolecular interactions can thereby be monitored on the single-molecule level with a resolution an order of magnitude better than what can be achieved by confocal microscopy. Multicolor detection makes a significant transition from simple colocalization of the different fluorescent spots to FRET once the two different fluorophores come close enough (<10 nm). A short-wavelength “donor” fluorophore being excited by light may transfer its energy nonradiatively through a dipole− dipole interaction to an “acceptor” fluorophore that emits at longer wavelengths. The efficiency of the energy transfer process (E) is a highly sensitive function of the distance (R) between the probes: E = [1 + (R/R0)6]−1, where R0, the socalled Förster radius, is a constant equivalent to the interprobe distance at E = 0.5. Under appropriate conditions, E can be extracted from the ratio of the fluorescence intensity of the donor to the fluorescence intensity of the acceptor. Because R0 assumes a value on the order of a few nanometers, which is comparable to the physical dimension of biological macromolecules, FRET serves as a spectroscopic ruler on the 3−10 nm scale, far below the diffraction limit or even the spatial resolution typically achieved in super-resolution or highresolution colocalization experiments. The observation of FRET between a single pair of fluorophores in 199673 represented a breakthrough for singlemolecule spectroscopy. smFRET methods have since been widely used to investigate the dynamic structure and interactions of proteins, nucleic acids, and macromolecule complexes at the nanoscale.8 The energy transfer efficiency also depends on the orientations of the donor and the acceptor. This effect limits the interpretation of FRET regarding interprobe distances. Long and flexible linkers between the biomolecule and the fluorescent probe are more likely to allow random orientation of the fluorophore that averages out the orientation dependence. However, even a long linker does not ensure mobility of the fluorophores, which might stick to the surface of the biomolecule. Another challenge is to distinguish the scenarios where low FRET results from large donor−acceptor distances from the cases where the acceptor is simply absent. A solution to this problem is the concept of alternating laser excitation (ALEX) scheme that alternately excites the donor and the acceptor.74 Donor excitation yields two observables, donor emission (D) and FRET (F), which are measured at the same time in two detectors. Acceptor excitation yields the observable acceptor emission (A), which is measured in the same detector as (F). The three observables are combined by calculating two ratios, E and S (eqs 2 and 3). The FRET efficiency E is dependent on the donor−acceptor distance. The stoichiometry ratio S is independent of the distance, and it reports the presence or the absence of the donor and acceptor even without proximity between the probes. E = F/(γ D + F)

S = (γ D + F)/(γ D + F + c A)

(3)

Typically, only γ is mentioned in the literature, and c is usually varied by adjusting the excitation intensity for the A channel. ALEX has proven to be a general and useful platform for smFRET experiments. Two-dimensional histograms of S versus E allow for virtual single-molecule sorting, somewhat in analogy with flow cytometry.74 In a simple system with a high FRET (short interprobe distance) and a low FRET (high interprobe distance) state, four different populations are typically resolved: the two FRET states at a stoichiometry value of about 0.5, in addition to a population that is lacking the acceptor (donoronly state, S = 1) and a population that is lacking the donor (acceptor-only, S = 0). ALEX is compatible with various time scales ranging from nanoseconds to milliseconds,75 and has been used to investigate structural dynamics of several biological systems including DNA replication by DNA polymerase or translation by ribosome.76,77

3. CELL BIOLOGY AND BIOCHEMISTRY OF GPCRs 3.1. GPCRs as Important Drug Targets

The GPCR superfamily has almost 1000 members.78 The receptor superfamily is subdivided into five classes named after their representative member: rhodopsin (class A), secretin (class B), adhesion (originally class B), glutamate (class C), and f rizzled/taste2.79 The rhodopsin receptor family is the largest category, comprising about 700 members in humans.80 All GPCRs share the seven transmembrane (TM) helical domains as a common structural framework. The extracellular Nterminus, intracellular C-terminus, and the loops between transmembrane helices are much less conserved and play a critical role in defining ligand−receptor and receptor−G protein interactions.3,78 GPCRs translate extracellular chemical messengers to specific intracellular responses that eventually lead to large-scale physiological effects. Despite the shared structural architecture, GPCRs respond to a large repertoire of stimuli of divergent chemical structures, such as ions, small molecules, lipids, peptides, or proteins. Not surprisingly, GPCRs are linked to a wide range of pathologies, including HIV, cancer, cardiac malfunction, asthma, neurological and inflammatory diseases, and obesity. In 2006, Overington et al. reviewed the genefamily distribution of drug targets and pointed out that 27% of FDA-approved drugs target class A GPCRs. A more recent estimate of the number of licensed medicinal drugs targeting GPCRs is 36%.2 Nonetheless, discovering new small-molecule drugs remains challenging; in many cases it is unclear why structurally similar drug candidates can induce very different physiological responses, sometimes with undesirable off-target side effects. 3.2. Assembly of the GPCR Signaling Complex

GPCRs are exquisitely regulated signaling machines. The tertiary complex of GPCR, ligand, and downstream adapter molecules is also called GPCR “signalosome”.39 How ligands differentially modulate the stoichiometry and the sequence of events in the dynamic assembly of the GPCR signalosome serves as the microscopic basis for pharmacology. As the name suggests, the canonical GPCR signalosome involves direct coupling between the receptor and a G protein (Figure 9a). Binding of an activating ligand, that is, an agonist, to the extracellular side of a GPCR induces conformational changes on the intracellular side of the receptor.3,81 Unlike

(2) H

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described arrestin signaling pathway is coupled to the mitogenactivated protein kinases (MAPK) in the cytosol. Arrestin also activates the phosphatidylinositol-3 kinase (PI3K) pathway and transcriptional regulation. 3.3. Spectroscopic and Structural Studies on GPCR Activation

How to activate a receptor, or how to turn it off, has been one of the most appealing questions in the GPCR field. As the ligand and the G protein bind to the opposite sides of the receptor, this allosteric effect must be mediated by a conformational change of the seven transmembrane helices. The most insightful experiments were done primarily on two prototypical GPCRs, the photoreceptor rhodopsin in the visual system and the β2 adrenergic receptor (β2AR) in the sympathetic nervous system. The biochemistry of rhodopsin commenced as early as in the 19th century when Willy Kühne established rhodopsin as the chemical basis for visual function.87 While Jokichi Takamine isolated adrenaline in 1900,88,89 the biochemical and biophysical characterization of β2AR truly gained momentum in the 1980s thanks to the cloning of adrenergic receptors.90−92 Even before the cloning of adrenergic receptors, the functional analogy between the lightactivated signaling pathway in the rod outer segment and the adrenaline-stimulated signaling pathway in hormone-sensitive cells had been observed by Mark Bitensky and Lubert Stryer.93−95 The sequence and structural homology between photoreceptors and adrenergic receptors cemented the parallelism, which is now known as the classic G proteincoupled receptor pathway.96 Both spectroscopic and structural studies require a significant amount of purified samples and rich experience with biochemical reconstitution. The high abundance of rhodopsin in the retina makes it possible to purify the receptors on a routine basis from native tissues. The large-scale production of β2AR was made possible by optimizing the purification of Nand C-terminal dual-tagged proteins using tandem affinity purification in combination with ligand-affinity chromatography.97 Rhodopsin is a unique GPCR in the sense that its native ligand, 11-cis-retinal, is covalently linked to the receptor. It acts as an inverse agonist until it gets activated by light and isomerizes into the agonist all-trans-retinal. The photochemistry of 11-cis-retinal can be traced by UV−vis spectroscopy. Experiments conducted at low-temperature revealed the existence of intermediate states in the course of receptor activation. However, the absorption spectra alone have not been very informative for understanding the conformational changes. It was EPR experiments on rhodopsin that provided the first clue that GPCR activation involves the rigid-body relative movement of TM3 and TM6.98 When the cytoplasmic ends of TM3 and TM6 were held together by metal ion binding99 or by disulfide cross-linking,98 photoactivated rhodopsin failed to activate transducin. These findings demonstrated that the relative movement between TM3 and TM6 was essential for rhodopsin activation. β2AR utilizes diffusible ligands with varying affinities. The pharmacology of β2AR is well-characterized and a large number of ligands are available, which makes β2AR an excellent model system for understanding how ligands differentially modulate GPCR conformations. Fluorescence experiments on β2AR pointed to similar structural changes upon activation as observed for rhodopsin,100,101 and suggested that agonists

Figure 9. Recognition of GPCRs by different adapter proteins. (a) On the extracellular side an agonist ligand (L) binds a GPCR, which in turn binds G proteins, G protein-coupled receptor kinases (GRKs), or arrestins on the cytosolic side. (b) Comparison of the inactive state (cyan, PDB: 1GZM)108 and the active state (orange, PDB: 3CAP)110 of rhodopsin reveals the outward movement of TM6 upon activation, a mechanism that seems to be conserved among family A GPCRs. (c) The crystal structure (PDB: 3SN6) of active-state β2AR (yellow) interacting with Gs (α, cyan; β, magenta; γ, green).119 (d) The crystal structure (PDB: 4ZWJ) of rhodopsin (orange) in complex with arrestin (blue).114

many other important transmembrane receptors such as ligandgated ion channel receptors or enzyme-coupled receptors, GPCRs lack catalytic activity and require the heterotrimeric G protein (Gαβγ) to serve as an intermediate between the receptor and the effectors. GPCR activation leads to GDP− GTP exchange in the Gα subunit and subsequent decoupling of the Gα subunit from the Gβγ subunit. The Gα and Gβγ subunits may independently mediate downstream signal transduction by modulating the level of second messengers such as Ca2+ and cyclic adenosine monophosphate (cAMP). Gα is a weak GTPase whose activity is regulated by GTPase activating proteins. The hydrolysis of GTP to GDP deactivates Gα and eventually results in the reassociation of Gα with Gβγ.82 GPCRs also signal through G protein-independent pathways (see also Figure 9a).83 The C-terminal serine and threonine residues on the activated receptors can be phosphorylated by G protein-coupled receptor kinases (GRKs), resulting in different phosphorylation barcodes. The phosphorylated receptors, in turn, lead to differential recruitment of arrestin.84 The roles of arrestin are 2-fold. First, by sterically blocking the G proteinbinding site on the cytoplasmic surface of the receptor, arrestin prevents further G protein activation. Thus, arrestin attenuates GPCR signaling, a process called “desensitization”. Meanwhile, arrestin also acts as an adapter molecule in the clathrinmediated endocytosis of the receptor to rapidly reduce the receptor copy number on the cell surface. The internalized GPCRs are either directed from the endosomes to the lysosome for degradation or recycled to the cell surface for future use in a desensitized form.85 Second, arrestin is capable of signaling in a G protein-independent manner.81,86 The bestI

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Figure 10. Overview of dynamic changes in GPCRs. GPCRs and their binding partners, which include ligands, adapter proteins, or other GPCR molecules, are in a dynamic conformational equilibrium.

increasing with structures from more than 30 different GPCRs in the public domain.

and partial agonists stabilize distinct conformational states.102−104 The appreciation of this conformational complexity turned out to be ultimately important to the subsequent crystallographic experiments: the key to obtaining crystals of GPCRs was to identify conditions that should stabilize the receptors in a single state. The fluorescence experiments also showed that the agonists themselves were insufficient to stabilize the activated receptor but additionally required a G protein. Therefore, additional engineering of the receptors, like constitutively activating mutations or G protein mimics, would be necessary for obtaining the active-state structure. The spectroscopic studies on GPCRs will be revisited in section 4.11.2. The structural biology of GPCRs began in 1993 when Schertler et al. presented a projection structure of rhodopsin at 7−9 Å resolution.105 In 2000, Palczewski et al. published the first high-resolution X-ray crystal structure of rhodopsin,106 and more refined structures ensued within a few years.107−109 The follow-up efforts were devoted to capturing the functionally relevant conformations of rhodopsin, with notable examples like the crystal structures of ligand-free opsin,110 opsin in its G protein-interacting form,111 and Meta-II rhodopsin.112,113 Thus, rhodopsin became the first GPCR whose inactive, active, and discrete intermediate states have been crystallized. In 2015, the long-awaited structure of the rhodopsin−arrestin complex was made possible by femtosecond X-ray crystallography.114 In 2007, the Kobilka group and the Stevens group reported the high-resolution structures of engineered β2AR in the inactive state.115−117 In 2011, the Kobilka group presented the structure of β2AR in its active state,118 and most extraordinarily, the high-resolution model of an active GPCR in complex with its G protein partner.119 This receptor−G protein complex structure reveals important contacts on the receptor−G protein interface and conformational changes that lead to the opening of the nucleotide-binding pocket. These data together have provided a framework for understanding the molecular basis of GPCR activation.120,121 It is remarkable how these structural studies corroborated the earlier spectroscopic data with precision and clarity. The active state structures of rhodopsin and β2AR both exhibit a marked outward shift of TM6. For β2AR, this conformational change is as large as 14 Å (Figure 9b).119 Further comparisons with an adenosine receptor,122−124 a muscarinic acetylcholine receptor,125,126 and an opioid receptor127,128 revealed a conserved molecular mechanism for GPCR activation. The structures are poised to accelerate drug development by providing threedimensional information on the ligand-binding pocket.129−131 Currently, the number of GPCR crystal structures is still rapidly

3.4. Conformational Diversity of GPCRs

The spectroscopic and structural studies on GPCRs rectified the simplistic two-state model for receptor activation, which assumed that there were merely the fully inactive state and the fully active state in equilibrium. Instead, GPCRs are better conceptualized as highly dynamic proteins (Figure 10) that sample an ensemble of interchanging conformations (Figure 11)

Figure 11. Conformational ensembles of GPCRs. Left panel: Schematic representation of theoretical conformational ensembles. Each rectangle defines the total conformation space of a GPCR. A given GPCR with no ligand exists in a resting ensemble of conformations (green). This conformational ensemble can be differentially modulated by ligand binding (blue and red). Each conformational ensemble may or may not contain conformations, which are recognized by each of the conformation-sensor proteins (dashed gray ovals). In this example, ligand 1 (blue) is a balanced ligand, whereas ligand 2 (red) is highly biased toward G protein activation. Such two-dimensional representation is, of course, an oversimplified picture of a highly multidimensional situation. Right panel: Arrestin recruitment versus G protein signaling for ligands of the same receptor. Balanced ligands define a diagonal axis, whereas biased ligands, which favor one pathway over another, are found off this axis.

The dynamical equilibrium within the conformational ensemble can be described by an energy landscape.132−136 The depths of the potential wells dictate the statistical distribution of each conformational state. The energy barriers correlate with the rates of exchange between states. Ligands act by modifying the shape of the energy landscape. Ligand-free receptors are mostly populating the inactive conformational states that do not recruit adapter proteins, with occasional exploration of active conformations giving rise to basal signaling J

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Figure 12. Fluorescent labeling schemes for GPCRs. (a) A fluorescently labeled antibody recognizing a conformational epitope. (b) A GFPnanobody as a conformational sensor for the activated receptor.198 (c) A fluorescent ligand specifically binding to the receptor. (d) Ligand-directed labeling based on Zn-aspartate coordination.270 (e) Intramolecular FRET as reporter of GPCR activation.231 (f) Intermolecular BRET as reporter of arrestin recruitment.245 (g) Combining a FlAsH-tag and a fluorescent protein for monitoring receptor activation.266 The smaller FlAsH, unlike CFP, does not disrupt the interaction between the receptor and the heterotrimeric G protein. (h) An N-terminal self-labeling protein tag. (i) Combination of a fluorescent ligand and a self-labeling tag (e.g., SNAP-tag) as a FRET pair for monitoring receptor−ligand binding.210 (j) Site-specifically labeled receptor. As compared to (d)−(i), the receptor is minimally modified. (k) FRET between a site-specifically labeled receptor and a fluorescent ligand.512 (l) A receptor carrying a fluorophore at the IC surface interacting with a fluorescent G protein.474

levels.137 The activation process should be viewed not as the switch from one inactive conformation to the active state, but as the shift of a whole ensemble of receptor conformations to another set of conformations that, on average, have a higher probability in recruiting adapter proteins.129,137,138 Comparing the spectroscopic and structural data of rhodopsin and β2AR revealed two distinct “personalities” of GPCRs.139 Dark-state rhodopsin is likely to represent the most energetically stable conformation of GPCRs whose activation involves the accumulative local structural changes that culminate in pronounced helix movement.121 The intermediate states on the activation trajectory of rhodopsin are short-lived, which suits its role as a photoreceptor. Therefore, the energy landscape of the inactive dark-state of rhodopsin features one predominantly populated potential well. By comparison, β2AR displays a greater conformational diversity, suggesting that its energy landscape, even in the unliganded basal state, is characterized by multiple populated potential wells. The concept of conformational diversity of GPCRs has profound therapeutic implications. GPCR ligands may activate

the receptor (agonists), block the receptor (antagonists), or silence the receptor (inverse agonists). Ligands that modulate the binding and function of the agonist are called allosteric modulators. Positive allosteric modulators (PAMs) enhance the agonist activity, negative allosteric modulators (NAMs) suppress the agonist activity, and silent allosteric modulators (SAMs) have neutral effects on agonist activity. A GPCR agonist is called a “biased” ligand, if it preferably elicits either G protein signaling, or arrestin signaling, or other noncanonical pathways. In other words, a biased ligand is more effective than a “balanced” ligand in shifting the ensemble to toward a particular state that engages one out of the many possible adapter proteins (Figure 11). This biased signaling paradigm can guide drug design: a ligand that selectively activates a pathway can avoid the undesirable physiological responses associated with the promiscuous activation of irrelevant pathways, while a ligand that selectively inhibits a pathway can ameliorate the disruption of the normal signaling.140−142 K

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dots171 have received increasing attention. As extrinsic fluorescent probes vary greatly in term of size and composition, attaching them to GPCRs requires specialized strategies according to their individual chemical properties. Here, we primarily review fluorescent labeling strategies for GPCRs (Figure 12).

3.5. Membrane Dynamics and Oligomerization of GPCRs

Research in the past 20 years has led to a firm appreciation of the crucial roles that oligomerization plays in regulating the function of various membrane receptors, such as protein tyrosine kinase receptors, cytokine receptors, TNF receptors, antigen receptors, etc.143 However, GPCRs have been classically described as self-sufficient monomeric receptors that form a ternary complex with a ligand and a G protein. This view is supported by the observations that monomeric rhodopsin and β2AR reconstituted into a high-density lipoprotein particle were capable of activating G protein.144−146 Monomeric rhodopsin solubilized in detergent micelles was also found to activate G protein at the diffusion limit.147 Nonetheless, the occurrence of GPCR homo- or heterooligomerization has been substantiated by mounting evidence. The oligomerization of heterologously expressed GPCRs has been extensively documented.148−151 Oligomerization of purified receptor reconstituted into lipid vesicles has also been reported.152−154 In several cases, GPCR oligomerization has been directly observed in native tissues. Atomic force spectroscopy revealed that the rhodopsin molecules assemble into higher orders in the native disc membrane.155,156 Fluorescence techniques have demonstrated the oligomerization of β-adrenergic receptors on cardiac myocytes157,158 and oxytocin receptor on mammary gland.159 The physiological relevance of oligomerization appears to be receptor-specific and remains under debate.160,161 It appears that GPCR oligomerization depends on the composition, thickness, and curvature of its membrane environment.152,162 For some receptors, oligomerization is a prerequisite for surface expression.163−165 Dimerization of GPCRs has also been implicated as a regulatory mechanism for ligand binding, G protein activation, and arrestin recruitment. In this model, the stoichiometry of ligand, receptor, and G protein (or arrestin) in the tertiary signaling complex can deviate from the classic 1:1:1 ratio.166 Promotion or inhibition of oligomerization by ligands has frequently been reported, implying the possibility of targeting the dimerization interface for therapeutic purposes.161,167 Overall, the physiological roles and regulation of GPCR oligomerization are not fully understood.

4.2. How to Specifically Target a GPCR

Specificity for targeting a GPCR or any protein of interest (POI) may be achieved based on one or a combination of the following principles. First, the POI can be targeted with specific reagents like ligands or antibodies. The specific affinity between the label and the POI arises from a series of weak, noncovalent interactions. Second, the recombinant DNA technology has made it routine to modify the nucleotide sequence to encode genetically a polypeptide tag into the POI. The tag and the original sequence are linked together through the amide bond. Third, certain chemical functionalities in the POI can be exploited to form a stable linkage through coordination or covalent bonding. Labeling strategies based on the molecular recognition of the receptor and the ligand, or of the receptor and the antibody, involve less engineering of the receptor. However, generating such specific reagents with high affinities can be laborious, if possible at all. The knowledge gained from a particular POI is not always applicable to other proteins. The genetically encoded tag can be a short peptide or a protein tag that folds into a tertiary structure. The POI and the tag should be fused in a way that they exist as individual modules without a major steric clash. Therefore, the targeted region in the POI needs to exhibit sufficient structural flexibility to tolerate the modification. The structural hallmark of GPCRs is the seven transmembrane helices that are connected with extracellular or intracellular loops. Even before the disclosure of high-resolution structures, it had been learned through the creation of functional chimeric α- and β-adrenergic receptors that the loops of the heptahelical receptors are able to accommodate major modifications.172 This insight later facilitated the crystallization of adrenergic receptors.115 The flexible loops, together with the N- and C-terminus of GPCRs, provided the possibility of extensive protein engineering. The fusion protein approach, despite numerous successful demonstrations, is inherently limited by the sheer size of the protein tag: the possibility of altering the native behaviors of receptors cannot be excluded a priori. Moreover, the compact TM region cannot be targeted. Strategies for selective labeling of some chemical functionality in the POI attempt to limit the modification to one or few residues. Such limited modification is difficult to achieve in a site-specific manner by targeting natural functionalities, as very often all amino acids occur more than once in a protein as large as a GPCR. In response to this challenge, the chemical biology community has resorted to bioorthogonal chemistries,173,174 that is, the reactions targeting functionalities that are not naturally present in living systems and that do not interfere with the native biological process. The bioorthogonal labeling strategies comprise two steps. First, a bioorthogonal reactive handle is introduced into the POI by unnatural amino acid mutagenesis or chemoenzymatic labeling. Second, a probe carrying a cognate reactive partner “clicks” with the bioorthogonal handle to form a permanent linkage. The ideal labeling methodology should be applicable in a general fashion to a wide range of POIs, minimally perturb their

4. LABELING OF GPCRs WITH BIOPHYSICAL PROBES 4.1. Overview

Biophysical studies of GPCRs critically rely on the ability to introduce a reporter probe precisely at the desired location without perturbing receptor function. When designing such experiments, three interrelated issues need to be taken into consideration: what kind of probes should be introduced, which region of the receptor needs to be modified, and how the probe can be anchored to the targeted region. The particular spectroscopic method being employed dictates the choice of the probe. Among the spectroscopic methods discussed above, IR, NMR, EPR, and MS are essentially in vitro methods; that is, the protein samples need to be isolated from the cells before characterization, where the probes are typically only one or a few atoms large. Fluorescence represents a more versatile tool, as it is widely applied both in vitro and in vivo with good temporal and spatial resolution. Accordingly, a wealth of fluorescent probes has been developed for the rapidly evolving fluorescence techniques. Fluorescent proteins168 and organic dyes169 have been the most popular choices, but in recent years lanthanide ions170 and quantum L

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intrinsic functionality, and allow for flexibility in choosing which region to target. However, these goals are rarely satisfied at the same time. When it comes to GPCRs, several additional aspects need to be taken into account. The first key consideration is the lifetime of the receptor. For the purified receptor, the half lifetime of denaturation typically ranges from hours to days depending on the stabilizing effect of the ligand and the type and ratio of the detergents and lipids. For GPCRs on the cell surface, their surface residence time may range from minutes to hours.175 The kinetics of the labeling reaction should be at least 1 order of magnitude faster than the lifetime of the receptor to ensure that the receptor remains functional. Second, the reaction kinetics determines the labeling stoichiometry. If the functional assay only involves individual receptors, substoichiometric labeling does not necessarily matter. However, in studies of ligand−receptor interaction or of receptor dimerization, the presence of unlabeled receptor complicates the analysis. Third, GPCRs are embedded in a lipidic environment. The reactions for labeling GPCRs should proceed under mild conditions and be compatible with the detergents and lipids in the system. Highly hydrophobic labeling reagents can be difficult to remove from the system, resulting in a high background.

the μ-opioid receptor (μOR),191 and the serotonin receptor.192 Such antibodies are powerful research and therapeutic tools.193 For example, the monoclonal antibody 2D7 recognizing the folded CCR5 receptor was used to screen for the optimal buffer composition to stabilize the purified receptor194 as well as to inhibit HIV viral entry.190 However, raising antibodies against a particular conformation of a receptor is challenging. Obtaining homogeneous receptor stabilized in a relevant conformation is far from being straightforward. Moreover, GPCRs are embedded in a lipid bilayer so that a significant part of the receptor is not accessible to the antibody. Hence, it is easier to develop conformation-sensitive antibodies for class B and class C GPCRs that possess a large and modular N-terminal domain rather than for class A GPCRs with a shorter N-terminal tail. 4.3.2. Nanobodies. In recent years, nanobodies, the singlechain monovalent antibody fragments engineered from heavychain antibodies of camelidae, became well-known in the GPCR field thanks to their crucial role in crystallizing the active-state β2AR.118,195,196 In addition to their high affinity and specificity, nanobodies (15 kDa) are much smaller than antibodies (150 kDa) and bear simpler post-translational modifications, which render them to be more water-soluble, diffusible, thermally stable, and better expressed in heterologous host cells. Not surprisingly, nanobodies emerged as promising tools for numerous laboratory and clinical applications. Immuno-imaging with nanobodies is under active exploration.197 The monovalence of fluorescently labeled nanobodies can be an advantage in probing receptor homodimerization. The easiness of heterologous expression makes it possible to engineer nanobodies into intracellular biosensors for the conformational change of GPCRs. GFP-tagged Nb80, a nanobody specific to the activated β2AR, allowed detection of receptor activation in endosomes (Figure 12b),198 which, otherwise, would be difficult to monitor by secondary messenger-based assays.

4.3. Immunofluorescence Using Antibodies and Nanobodies

4.3.1. Antibodies. Historically, dating back to the seminal work of Albert Coons in the 1940s and 1950s,176 the method of choice for labeling GPCRs on the cell surface has been immunofluorescence based on antigen−antibody reactions. In the prerecombinant DNA era, antibodies were raised against only a few GPCRs, like rhodopsin and β-adrenergic receptors, which were available in sufficient quantities from biochemical purification.177,178 However, not all antibodies simply bind with the receptor in a passive way; sometimes they modulate receptor signaling and its oligomerization state.179−181 Therefore, antibodies raised against receptors may not be the perfect imaging tools. The introduction of recombinant DNA technology in the 1970s made it possible to tag the POI with a foreign epitope that allows recognition by specific antibodies independent of the POI.182 A wealth of such epitopes and specific antibodies has been made available to the scientific community.183 These reagents have been used to detect GPCRs in a variety of applications, such as Western blot, flow cytometry, or immunofluorescence. As compared to radioactive labeling,184 immunofluorescence represents an environmentally safe technique for determining the cellular localization of GPCRs, and the spatial resolution is significantly better.185,186 ImmunoFRET was used in the early experiments that directly showed GPCR dimerization on the cell surface. The energy transfer between fluorescently labeled antibodies specific to individual monomer demonstrated the physical proximity of receptors.187−189 Although it is not difficult to derivatize antibodies with fluorophores, immunofluorescence is limited by the properties of antibodies, which are bivalent, nonspecific in some cases, and unable to penetrate the plasma membrane of living cells. Conformation-specific antibodies, on the other hand, selectively recognize a particular folded state of the receptor by interacting with its tertiary structural motif (Figure 12a). Conformation-sensitive antibodies have been reported for only a few receptors, for example, the chemokine CCR5 receptor,190

4.4. Fluorescent Ligands

Synthetic ligands have always played an important role in understanding GPCRs. In fact, the very presence of adrenergic receptors on the cell membrane was demonstrated by radioactive high-affinity ligands.199,200 The molecular recognition between GPCRs and their ligands can be exploited to label the receptors. Like conformation-sensitive antibodies, ligands selectively bind with correctly folded, functional receptors, thus excluding the interference of denatured receptor. However, ligands may also alter the behavior of the receptor, for instance, inducing its internalization.85 Fluorescently labeled high-affinity ligands have been used to quantify the amount of functional receptors in vitro,201 or to visualize receptor expression on the cell surface (Figure 12c).202,203 Monitoring the FRET signal between fluorescent ligands and fluorescently labeled GPCRs enables direct visualization of ligand binding events in vitro or in living cells.204 Typically, the receptors need to be tagged at their extracellular surface to achieve efficient energy transfer.205 Previously, the FRET and BRET experiments using fluorescent protein/luciferase-tagged GPCR constructs (described in greater details in sections 4.6 and 4.7) report binding of fluorescent protein-tagged binding partners (G protein subunits, arrestins, other GPCRs, etc.). Such experiments allow indirect detection of ligand binding as inferred from binding with the adapter proteins. In contrast, monitoring binding of fluorescent ligands reports ligand binding events that are not necessarily followed by the binding M

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of the adapter proteins. Single-molecule fluorescence detection of the dynamic assembly of the ligand−receptor-G protein ternary complex is under exploration.206 Fluorescent ligands have also been used to study GPCR oligomerization. To date, most of the energy transfer studies on GPCR oligomerization have been performed on transfected cells overexpressing the receptors, especially for BRET experiment in which the expression is often maximized to enhance the signal. The physiological relevance of these approaches has been under debate.207,208 Although overexpression does not necessarily lead to artifacts that would invalidate the functional relevance of observed oligomerization, it would be ideal to detect wild-type receptor oligomerization directly in native tissues. Time-resolved FRET between fluorescent ligands, thanks to its sensitivity and specificity, has provided a possible solution. In an example, europium and Alexa Fluor 647 were attached to high-affinity peptide ligands for the oxytocin receptor, whose oligomerization was detected both in transfected cells and in the rat mammary gland.159,209 With an increasing list of fluorescent ligands and lanthanidelabeled GPCRs at hand, high-throughput time-resolved FRET assays210,211 were developed to screen for drug candidates212 or to validate the functionality of engineered receptors.213 Fluorescently labeled ligands also enable single-molecule studies of GPCRs. For example, Alexa Fluor 647-labeled MIP1α was used to selectively visualize CCR5 chemokine receptors captured on a TIRF surface.39 Single-molecule tracking experiments using fluorescent ligands will be discussed later in section 5.1. Another tool to study GPCR dimers is a fluorescent bivalent ligand. The cooperativity between two pharmacophores gives rise to a high and specific affinity of the bivalent ligand for the receptor dimers.214 The key is to tweak the length and composition of the linker so that bivalent ligands exhibit the expected subtype selectivity and lipophilicity. The reports on fluorescent bivalent ligands, however, have been rare. In one example, a fluorescent bivalent ligand for the chemokine CXCR4 receptor was used to visualize its dimerization in transfected cells.215 Preparing fluorescently labeled ligands and validating their affinities and efficacies are not straightforward. Adding a synthetic dye to a ligand may drastically alter its pharmacology, especially for small-molecule ligands. Fluorophore-derivatized peptide ligands are relatively easy to prepare because a linker can be introduced to separate the dye from the pharmacophores. There are a few reviews on the progress in developing fluorescent ligands for GPCRs.193,202,216−218

Figure 13. Ligands that form covalent linkages with GPCRs. (a) BABC, a carazolol analog that reacts with a nucleophilic residue on β2AR.220 (b) FAUC50, which forms a disulfide with an engineered cysteine.221 (c) Labeling reagent for ligand-directed tosyl (LDT) chemistry. (d) Labeling reagent for ligand-directed acyl imidazole (LDAI) chemistry. Both (c) and (d) react nonspecifically with a proximal nucleophile and do not require a priori modification of the POI.

moiety.224 When the ligand binds to the receptor, a proximal nucleophilic residue on the POI displaces the ligand through an SN2 reaction to form a covalent linkage with the probe. LDAI chemistry was reported to have higher labeling efficiencies than LDT224 and has been demonstrated for various membrane proteins, including the GPCR bradykinin receptor.225 Liganddirected labeling strategies avoid coexpression of post-translational modification enzymes or generation of a fusion protein. Notably, LDT and LDAI chemistries are among the few methods available for labeling endogenous membrane proteins. 4.6. Fluorescent Proteins and Förster Resonance Energy Transfer (FRET)

The seminal work of Roger Tsien and Martin Chalfie226−228 has popularized fluorescent proteins (FPs), such as the green fluorescent protein (GFP), as an essential tool for studying protein function in cells and tissues.229 How to choose a suitable fluorescent protein for specific questions has been discussed elsewhere.168 Fluorescent proteins opened an avenue for interrogating the behaviors of GPCRs in live cells and animals. The first example of an FP-tagged GPCR was the prototypic β2AR that carried a GFP at its C-terminus.230 Later reports showed that modifications at the N-terminus,205and the third IC loop231 also yielded functional receptors. FP-tagged GPCR constructs quickly became a standard approach for evaluating the dynamic localization and trafficking of GPCRs.232,233 A central question in the GPCR field is the molecular details of receptor activation. In vitro fluorescence experiments on purified β2AR showed that receptor activation occurred on a time scale of minutes, which was significantly slower than the activation in cells.100,102,234 This inconsistency posed a need for developing cell-based assays for receptor activation. Vilardaga et al. generated parathyroid hormone receptor and α2A adrenergic

4.5. Ligand-Directed Labeling

The ligand-directed labeling strategy harnesses the specific binding between a receptor and a ligand to create a covalent linkage through a proximity-induced reaction. The ligand carries an electrophilic reactive moiety that reacts with a nucleophilic residue, such as cysteine, tyrosine, or lysine, situated on the extracellular surface close to the binding pocket, in a position favoring modification.219 Such ligands have been exploited to deduce the structure of β2AR and facilitate its crystallization (Figure 13a,b).220−222 The Hamachi group developed ligand-directed tosyl (LDT) chemistry and ligand-directed acyl imidazole (LDAI) chemistry with the aim of achieving fluorescent labeling (Figure 13c,d). The ligand and the probe are connected with a phenyl sulfonate moiety (LDT)223 or an alkyl-oxy acyl imidazole (LDAI) N

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Figure 14. Peptide tag-based fluorescent probes. (a) FlAsH (fluorescein arsenical hairpin binder).262 (b) ReAsH (resorufin arsenical hairpin binder). Both FlAsH and ReAsH bind to a genetically encoded tetracysteine motif. (c) RhoBo (rhodamine-derived bisboronic acid) that binds to a tetraserine motif.268 (d) Zn(II)-DpaTyr-tag. The four Zn(II) atoms coordinate with the oligo-aspartate tag fused to the N-terminus of the receptor, facilitating the formation of a thioester bond with the N-terminal cysteine.271 (e) Template-directed labeling based on a coiled-coil motif.272

receptor (α2AAR) constructs carrying the cyan fluorescent protein (CFP) in the third IC loop and the yellow fluorescent protein (YFP) at the C-terminus. The FRET between CFP and YFP served as a reporter for the cytoplasmic conformational change (Figure 12e). On the basis of the FRET data, it was found that the time scale of GPCR activation in living cells varied among receptors, from milliseconds for α2AAR to seconds for parathyroid hormone receptor.231 These values are, nonetheless, 1 or 2 orders of magnitude smaller than the activation kinetics measured for β2AR in vitro, demonstrating the physiological relevance of cell-based assays. Another popular application of FP-tagged GPCRs is to assess the oligomerization of receptors by monitoring the FRET.148,149,235 As compared to immunofluorescence, this strategy eliminates the need for optimizing the staining protocols and avoids the use of bivalent antibodies. GFP (27 kDa) is only modestly smaller than class A GPCRs (typically 40−60 kDa). Whether adding a bulky fluorescent protein to the receptor interferes with its native behavior requires a case-by-case evaluation. The early reports on Cterminally GFP-tagged versions of β2AR,230 cholecystokinin receptor type A,236 and cAMP receptor237 described that the labeled receptors resembled the wild-type receptors in terms of ligand binding, localization, trafficking, and downstream signaling. Contrary to these observations, the Milligan group reported that the additional C-terminal fluorescent protein affected the rate of adrenergic receptor internalization.238 As the IC loops are required to engage the heterotrimeric G protein,119,239 inserting FPs into the third IC loop was more problematic than fusing it to the termini.231

Protein engineering has not only produced a palette of fluorescent proteins spanning the visible wavelengths, but also yielded fluorescent sensors for protein conformational change, such as circular permuted GFP (cpGFP),240 or for protein− protein interaction, such as split GFP.241 These FP variants can be encoded into a single polypeptide, and the detection is independent of FRET. To our knowledge, no application of cpGFP to GPCRs has been reported. Recently, Jiang et al. described a GPCR construct tagged by split GFP on the extracellular surface.242 Split GFP consists of two complementary fragments that spontaneously assemble into a complete fluorescent β barrel. The smaller fragment (β strands 10 and 11) could be inserted into the third extracellular loop of two functional GPCRs, and that the larger fragment (β strands 1−9) was supplied exogenously to give cell-surface labeling. It can be envisioned that split GFP may serve as an alternate detection scheme for probing receptor oligomerization. 4.7. Luciferase and Bioluminescence Resonance Energy Transfer (BRET)

Bioluminescence resonance energy transfer exploits the nonradioactive energy transfer between the luminescence of Renilla luciferase (RLuc, donor) and a fluorescent protein (FP, acceptor) to detect intermolecular interactions.243,244 BRET does not require extrinsic excitation, thus bypassing the some unfavorable aspects of FRET, photobleaching, cross-excitation of the acceptor, background autofluorescence, and cellular damage due to high-power illumination. The ratiometric nature of BRET enables quantitative measurement of protein−protein interaction with good reproducibility. O

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Figure 15. Self-labeling protein tags. (a,b) Both SNAP- and CLIP-tag derive from O6-methylguanine-DNA methyltransferase with C145 as the active site.276,279 (c) The Halo-tag derives from haloalkane dehalogenase whose active site D106 forms an ester bond with the chloroalkane linker.300 (d) The TMP-tag noncovalently binds with trimethoprim and brings the α,β-unsaturated carbonyl (i)310 or sulfonyl (ii)311 into proximity of the engineered reactive cysteine L28C.

4.8. Peptide-Based Tags

Similarly to FRET, BRET has been used to examine receptor oligomerization.245−248 BRET and FRET using tagged GPCR constructs provided a large body of evidence, suggesting receptor oligomerization as a fundamental aspect of GPCR regulation.148,149,249,250 BRET can be used as a reporter for arrestin recruitment. In such an experiment, the receptor carries RLuc in its C-terminus, and arrestin fused with a suitable FP (Figure 12f).245,251 Because only activated GPCRs bind with arrestin, this BRET scheme was used to screen for GPCR agonists.252−254 The sandwich construct of β-arrestin tagged with the BRET pair served as an arrestin-specific conformational sensor255 and showed that arrestin adopted heterogeneous conformations in response to different ligands.256 Two recent studies further demonstrated that GPCRs impose distinctive arrestin conformations reflecting the stability of the receptor−arrestin complex.257,258 A BRET assay for G protein activation was also developed,259 which showed that kinetics of receptor-G protein binding was similar to the kinetics of receptor conformational change as measured by intramolecular FRET.231 The capability of quantifying arrestin recruitment and G protein activation provided evidence for biased agonism of GPCRs.140,260 For example, Masri et al. employed BRET to compare arrestin recruitment and G protein activation for the D2 dopamine receptor, and found that various antipsychotics shared the feature of antagonizing arrestin recruitment, but produced different effects in G protein activation.261 These findings are instructive for drug design.

4.8.1. Arsenical Hairpin Binders Specific for the Tetracysteine Tag. Fluorescein arsenical hairpin binder (FlAsH) tag technology harnesses the coordination between sulfur and arsenic to chelate a biarsenical derivative of fluorescein to a tetracysteine peptide motif genetically encoded into the POI (Figure 14a,b).262 The variants of suitable tetracysteine motifs range from a short version (CCPGCC)263 to longer and more specific versions (FLNCCPGCCMEP or HRWCCPGCCKTF),264 and split cysteine pairs spread over tertiary contacts in proteins. The green FlAsH and the redshifted variant ReAsH (resorufin arsenical hairpin binder) are fluorogenic reagents that are provided as nonfluorescent complexes with ethanedithiol. They become fluorescent only upon binding with the tetracysteine motif. The membrane permeability of FlAsH and ReAsH makes them suited for labeling the IC surface of membrane proteins or cytoplasmic proteins. FlAsH labeling is fast, stable for hours, and reversible by adding dithiols. Optimizing the labeling protocol reduces the potential cytotoxicity and nonspecific labeling.265 Hoffmann et al. presented a successful demonstration on how to use the CFP/FlAsH FRET pair to monitor the cytoplasmic conformational changes of GPCRs (Figure 12g).266 The FlAsH tag proved a superior FRET acceptor than YFP: its reduced size conferred greater sensitivity to small conformational changes and caused less alteration to the G proteininteracting surface. As compared to YFP/CFP tagged constructs, the CFP/FlAsH tagged adenosine A2A receptor (A2AR) exhibited similar activation kinetics but substantially higher adenylyl cyclase stimulating activity. In a subsequent P

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study, Vilardaga et al. showed that morphine binding to μOR suppressed the Gi signaling of α2A adrenergic receptor (α2AAR), as was monitored through the CFP/FlAsH FRET signal, providing new insights into the functional relevance of receptor dimerization.267 4.8.2. Bisboronic Probe Specific for the Tetraserine Tag. Rhodamine-based bisboronic acid (RhoBo) tag, being conceptually similar to FlAsH/ReAsH, is based on the coordination of boron and the hydroxyl groups in a tetraserine motif (Figure 14c).268 Originally designed for detecting the hydroxyl-rich monosaccharide,269 RhoBo tag turned out to prefer the tetraserine motif by 4 orders of magnitude. This RhoBo tag reagent is fluorogenic, cell-permeable, and nontoxic for cells. However, it suffers from the off-target labeling of endogenous proteins with tetraserine-like sequences. To our knowledge, there is no report on labeling GPCRs with RhoBo tag. 4.8.3. Tetranuclear Zinc(II) Probe Specific for the Oligo-aspartate Tag. The Hamachi group developed a motif/probe pair comprised of a cysteine-containing oligoaspartate tag and a tetranuclear Zn(II) probe with a reactive αchloroacetyl moiety (Figure 14d).270 The oligo-aspartate tag was fused to the N-terminus of the POI. The coordination between aspartate and Zn(II) facilitates the nucleophilic attack of a cysteine thiol on the chloroacetyl moiety, resulting in a stable N-terminal tagged protein. The tetranuclear Zn(II) probe was used to visualize the internalization of the bradykinin receptor.271 4.8.4. Template-Directed Labeling Based on a CoiledCoil Motif. In addition to the coordination of metal ions and amino acids, it is also possible to utilize peptide secondary structure motifs as recognition motifs. For example, two sizematched coiled-coil peptides were anchored to the human neuropeptide Y2 receptor and to a fluorophore, respectively. The coiled-coil structure facilitates the transfer of the fluorophore to the tagged receptor through an acyl transfer reaction (Figure 14e).272 The neuropeptide receptors and dopamine receptor labeled through this method retained the affinity for ligands and the ability for internalization and recycling.273 Template-directed labeling based on the coiledcoil motif only targets the extracellular surface of membrane proteins but gives permanent labeling.

(Figure 15a).276 SNAP-tag can be fused to POIs and then be labeled with synthetic dyes linked to benzylguanine.277,278 Subsequent engineering of O6-methylguanine-DNA methyltransferase yielded the CLIP-tag that accepts benzylcytosine derivatives (Figure 15b).279 The mutual orthogonality of the SNAP- and CLIP-tags enabled simultaneous labeling of multiple POIs in the same cellular context. The modular designs of benzylguanine and benzylcytosine substrates have enabled labeling of GPCRs with a broad spectrum of synthetic dyes,280,281 with a lanthanide,282 as well as with quantum dots.283 While SNAP- and CLIP-tags (19 kDa) are only slightly smaller than GFP (27 kDa), importantly, they offer greater freedom in choosing fluorescent reporters with desired photophysical properties.284 Similar to fluorescent proteins and fluorescently labeled antibodies, self-labeling protein tags are useful in real-time tracking of GPCR localization.285−287 A study based on the segregation of tagged proteins at cell division showed that even fluorescent proteins with monomerizing mutations suffered from some tendency to oligomerization, while the SNAP-tag caused the least perturbation to the localization of the labeled protein.288 This finding might suggest that the studies involving SNAP-tagged GPCRs are more likely to give an accurate account of receptor localization and oligomerization. In the GPCR field, SNAP-/CLIP-tag technology is notable for popularizing the application of time-resolved FRET techniques based on lanthanide emitters. The luminescence of lanthanide/macrocycle complexes, like terbium cryptate or europium cryptate, has a large Stokes shift as well as a significantly longer luminescence lifetime (∼milliseconds) than the intrinsic fluorescence arising from biomolecules (<10 ns). On the basis of these properties, lanthanide emitters can be paired with far-red fluorophores to achieve highly sensitive FRET measurements, in which a judicious choice of the measurement window suppresses the background signal (Figure 12i).289 Previously, lanthanide labels were typically conjugated to POIs through reactions with cysteines or lysines, which limited their utility to purified proteins.290 Lanthanidelabeled antibodies suffer from all of the inherent limitations of antibodies, and their preparation is time- and material-intensive. The now commercially available lanthanide-labeled SNAP/ CLIP substrates offer a standardized method for attaching lanthanide probes to GPCRs.210,282 Time-resolved FRET is useful in probing the binding events between SNAP-tagged GPCRs and fluorescently labeled ligands, which has been adapted to high-throughput format for screening receptor−ligand interaction.210,211 This assay is particularly suited for studying peptide ligands that can be modified with far-red fluorophores with relative ease. The power of time-resolved FRET has also been harnessed to assess the homo-oligomerization state of the class A metabotropic glutamate receptor and the class C γ-aminobutyric acid (GABA) receptor,282,291 and then quickly adopted to study chemokine receptors,292 β-adrenergic receptors,158 and the muscarinic acetylcholine receptor.293 An elegant single-molecule surface-tracking study for SNAPtagged GPCRs revealed different oligomerization levels of βadrenergic and GABAB receptors.294 In a single-molecule FRET study, the SNAP-tagged construct was used to understand the conformation dynamics of the metabotropic glutamate receptor dimers.295 The mutually orthogonal SNAP- and CLIP-tags were combined to probe the heterodimerization between different GPCRs, such as the cannabinoid and the orexin

4.9. Chemoenzymatic Labeling Based on Self-Labeling Protein Tags

Chemoenzymatic labeling methods exploit the exquisite molecular recognition mechanism between enzymes and substrates to create a specific covalent linkage between the label and a tag encoded into the POI.274 The earliest report on chemoenzymatic labeling of a GPCR dated back to 1978, when the Stryer group used transglutaminase to label the cytoplasmic surface of rhodopsin with synthetic dyes.275 However, transglutaminase-mediated labeling lacked site-specificity and quickly lost importance to chemical modifications of cysteines. 4.9.1. SNAP-Tag and CLIP-Tag. Modern chemoenzymatic labeling technologies harness the power of directed evolution technology to tailor the property of the enzymes. Currently, the most popular chemoenzymatic approach for labeling GPCRs is based on self-labeling proteins, which, as the name suggests, catalyze the covalent modification of themselves. The founding member of self-labeling proteins is the SNAP-tag derived from the mammalian O6-methylguanine-DNA methyltransferase that utilizes O6-benzylguanine (BG) derivatives as its substrate Q

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Figure 16. Chemoenzymatic labeling strategies using post-translational modification enzymes. (a) The engineered enzyme specifically transfers a probe to the acceptor sequence fused to the receptor. The probe contains a bioorthogonal reactive handle for subsequent labeling. The enzyme can be coexpressed in cells or supplied in vitro. (b) Biotin ligase-mediated labeling and the substrates: biotin (i), a ketone analogue (ii),312 an alkyne analogue (iii), and an azide analogue (iv).316 (c) Lipoic acid ligase-mediated labeling and the substrates: lipoic acid (v), an azide analogue (vi),318,540 a fluorinated aryl azide analogue for photo-cross-linking (vii),320 and a resorufin analogue for fluorescent imaging (viii).323 (d) Sortase-mediated labeling (Sortagging) that attaches a peptide tag to the POI. (e) Formylglycine generating enzyme-mediated oxidation resulting in a reactive aldehyde handle for subsequent labeling. (f) Ascorbate peroxidase (APEX) for proximity-dependent labeling of an electron-rich amino acid residue, e.g., tyrosine.

receptors,296 metabotropic glutamate receptor subunits,297 the dopamine D2 and the ghrelin receptors,298 the dopamine D2 and D3 receptors,299 etc. 4.9.2. Halo-Tag. Another member of the self-labeling protein family is the Halo-tag, a modified haloalkane

dehalogenase (33 kDa), whose active site aspartate forms an ester bond with a chloroalkane through an SN2 reaction (Figure 15c).300 Resin derivatized with Halo-tag substrate is useful for purifying the tagged POIs. There have been fewer reports on Halo-tagged GPCRs than on SNAP-/CLIP-tagged ones. A CR

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handles;318,319 (2) photo-cross-linking using fluorinated aryl azide;320 and (3) fluorescence imaging with hydroxycoumarin,321 Pacific Blue,322 and a red-shifted fluorophore resorufin.323 4.10.2. Sortase. The Ploegh group chose to engineer sortase, a transpeptidase isolated from Staphylococcus aureus or Streptococcus pyogenes, to specifically attach probes to a recognition sequence (Figure 16d).324 Sortase catalyzes the transpeptidation reaction for a conserved 5-amino-acid motif. First, sortase cleaves the amide bond between a threonine and a glycine in the protein to form an activated thioester intermediate. The addition of polyglycine-derivatized labels recreates the amide bond, resulting in a label directly linked to the carboxylic end of the recognition motif. As the active site of sortase does not have to accommodate the entire polyglycine substrate, there is no limit on the size and chemical property of the probe. Sortase-mediated labeling, or briefly termed as sortagging, has been demonstrated for several membrane proteins,324,325 including platelet-activating factor receptor, a class A GPCR.326,327 Theoretically, the 5-amino-acid recognition sequence for sortase can be genetically encoded at the Nterminus, the C-terminus, or the loops.328,329 However, in the course of transpeptidation, the POI is likely to disintegrate unless otherwise stabilized by a disulfide bond. By comparison, biotin ligase-mediated or lipoic acid ligase-mediated labeling is not restricted to the two termini of POIs, as both enzymes modify the side chain of the acceptor sequence rather than altering the primary structure. 4.10.3. Formylglycine-Generating Enzyme. The Bertozzi group developed a strategy based on the formylglycinegenerating enzyme (FGE), an enzyme responsible for creating the formlyglycine active site in sulfatases for hydrolyzing sulfate esters (Figure 16e).330 FGE oxidizes a conserved cysteine thiol in a 5-amino-acid motif into an aldehyde group that can be subsequently modified by ketone-reactive chemistries.331−333 4.10.4. Ascorbate Peroxidase. The Ting group tailored soybean ascorbate peroxidase (APEX) for proximity-dependent, promiscuous labeling (Figure 16 f).334−336 In the presence of hydrogen peroxide, APEX oxidizes phenols to generate short-lived phenoxyl radicals that covalently label proximal electron-rich amino acid residues, such as tyrosine, tryptophan, histidine, and cysteine, independent of any recognition sequence. The phenol substrate was derivatized with biotin or bioorthogonal reactive handles for subsequent detection. The 28-kDa APEX can be genetically fused to the POI and targeted to different cellular regions. This labeling scheme is particularly suited for proteomic profiling in enclosed cellular compartments, for example, mitochondria, which restrict the diffusion of the phenoxyl radicals.334,337 The oxidizing activity of APEX has also been harnessed to catalyze the formation of osmiophilic polymer in situ to give EM contrast.338 4.10.5. Applications of the Engineered Posttranslational Enzymes. The enzymatically attached biotin handle or bioorthogonal reactive handles provided an anchor site for subsequent attachment of fluorescent reporter and live-cell imaging.313,315,339,340 The bioorthogonal chemistries will be illustrated in greater detail in section 4.12.3. Coexpression of biotin ligase with GPCRs fused with the acceptor sequence in a mammalian system has yielded quantitatively biotinylated receptors in large scale,341,342 providing an alternate to His6tag purification. Chemoenzymatic labeling methods rely on the proximity between the post-translational modification enzyme and the target protein, which has been exploited by interaction-

terminal fusion with a Halo-tag was found to facilitate the bacterial expression and subsequent purification of functional cannabinoid receptor CB2.301 Halo-tagged GPCRs have been labeled with small-molecule fluorophores302−305 or quantum dots306 for single-particle tracking experiments in live cells. 4.9.3. TMP-Tag. A third self-labeling protein is the TMP-tag (18 kDa). The first-generation TMP-tag harnessed the highaffinity interaction between E. coli dihydrofolate reductase (eDHFR) and its small-molecule inhibitor trimethoprim (TMP) to form long-duration and yet reversible binding (Figure 15d).307,308 The second-generation TMP-tag exploited a proximity-induced reactivity to create a covalent linkage between the engineered active site cysteine and the label containing an α,β-unsaturated carbonyl moiety193,309,310 or a sulfonyl group.311 It is worth noting that eDHFR is an enzyme essential for tetrahydrofolate synthesis in bacteria, and TMP was originally developed as an antibiotic targeting folate pathway. Consequently, the binding mode between eDHFR and TMP has been clearly characterized. The development of TMP tag stands as a good example on how to take advantage of the historical legacy from another field. The self-labeling protein tags may be anchored to both the N- and the C-terminus of GPCRs. In the case of an N-terminal fusion, a signal peptide is often desirable to facilitate the trafficking of tagged receptors. As the labeling reagents need to be applied exogenously, the feasibility of C-terminally tagged receptors essentially depends on the membrane permeability of labeling substrates. A noteworthy advance for live-cell labeling is the development of fluorogenic substrates for self-labeling tags, which will be described in section 4.13. 4.10. Chemoenzymatic Labeling Based on Posttranslational Modification Enzymes

The size of self-labeling protein tags always raises questions regarding their effects on protein functionality. To circumvent this problem, a variation of chemoenzymatic labeling takes a different route: a short recognition peptide sequence is genetically encoded into the POI, which can be then specifically modified by a suitable posttranslational modification enzyme coexpressed in the same cellular context (Figure 16a). The substrate selectivity of the enzyme, together with its spatial proximity with the POI, ensures specific labeling. 4.10.1. Biotin Ligase and Lipoic Acid Ligase. The Ting group pioneered the approach of redirecting biotin ligase312 and lipoic acid ligase313,314 for protein labeling. The E. coli biotin ligase BirA catalyzes the formation of an amide bond between the carboxylic group of biotin and the ε-amino of a lysine situated in a 23-amino-acid recognition sequence (Figure 16b), also known as acceptor sequence. This 23-amino-acid acceptor sequence was further optimized to a 15-amino-acid version.312 Two orthogonal acceptor sequences were developed for E. coli312 and yeast315 biotin ligases to enable double labeling. The substrate repertoire was expanded to include ketone, azide, and alkyne biotin analogues.312,316 It was later discovered that a point mutation in BirA impaired the sequence specificity of the acceptor peptide, generating an enzyme that would promiscuously biotinylate a peptide substrate in proximity.317 The E. coli lipoic acid ligase attaches a lipoic acid to an optimized 13-amino-acid recognition sequence (Figure 16c).313,314 Lipoic acid ligase was engineered to incorporate probes for various purposes: (1) bioorthogonal labeling using aliphatic azide, aryl-aldehyde, or aryl-hydrazine as the reactive S

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Figure 17. Labeling the cysteines of GPCRs. (a) Popular cysteine chemistries (top to bottom): disulfide exchange using a disulfide reagent; alkylation using an alkyl halide, e.g., halide acetamide; Michael addition with α, β-unsaturated carbonyl compounds, e.g., maleimide. (b) The crystal structures of rhodopsin (Rho, PDB: 1U19) and β2AR (PDB: 2RH1) with the native cysteines highlighted (red, intracellular free cysteines; green, Cterminal palmitoylated cysteines; blue, transmembrane cysteines; orange, extracellular cysteines that form disulfide bond; yellow, extracellular free cysteines). Rhodopsin has 10 cysteines, and β2AR has 13 cysteines. C378 and C406 are reported to form disulfide bond during purification.102 The structure of β2AR lacks the C-terminus, so C378 and C406 are not shown. Palmitoylation of membrane proteins is a dynamic process.402 Thus, the supposedly palmitoylated cysteines are potentially available for modification. (c) Examples of biophysical probes attached to cysteine residues (from left to right): the spin label tetramethyl pyrrolidine-N-oxyl nitroxide (PROXYL) attached by disulfide exchange;98 PROXYL derivatized by iodoacetamide (IA-PROXYL);394 cysteine modified with monobromobimane (mBB, sometimes abbreviated as mBBr) as the smallest extrinsic fluorescent reporter;378,381,386 tetramethylrhodamine maleimide (TMR-ML) as an environmentally sensitive probe;103,104 2-bromo-4(trifluoromethyl)acetanilide (19F-BTFA) as an NMR probe;394,395 D5-N-ethylmaleimide (D5) as a MS probe.398,399

amino acids in cell culture (SILAC) should be used to improve the spatial resolution.336 Previously, GPCR in organelles has been less studied than the receptors expressed on cell surface. As compared to the plasma membrane, the organelle membranes are more difficult to purify and less amenable to the classic labeling techniques. Proximity-dependent labeling methods might be instrumental for interrogating GPCR signaling in organelles like endosomes345,346 and mitochondria.347 So far there have been few reports on applying proximity-dependent enzymatic labeling reactions to GPCRs. Nonetheless, the rapid expansion of this chemoenzymatic labeling toolkit is disposed to produce an impact on GPCR research.

dependent probe incorporation mediated by enzymes (IDPRIME)318,343 to probe the physical interactions between biomolecules. Two interacting proteins are individually tagged with either the enzyme or the acceptor sequence. The substrate specificity of the enzymes ensures good signal-to-noise ratio. This approach has been applied to assess the dimerization between GPCRs in the cellular context (Figure 26g).344 The receptors are tagged with either biotin ligase or the acceptor sequence. The more frequently oligomerization occurs, the higher is the probability that the acceptor sequence-tagged receptor becomes biotinylated. One limiting factor of this approach is that the biotin ligase and lipoic acid ligase are 35 and 38 kDa, respectively, which necessitates additional assays to validate the functionality of the fusion protein. Proximity-dependent labeling strategies using enzymes with promiscuous peptide substrates are powerful tools for identifying unknown protein−protein interaction. Roux et al. developed a method termed “BioID” that utilizes the promiscuous biotin ligase to tag neighboring proteins.317 APEX-based proximity labeling is conceptually similar to BioID. However, the diffusion of phenoxyl radicals might result in labeling of proteins several nanometers away. Therefore, detection schemes like stable isotope labeling by

4.11. Classic Approach for Site-Specific Labeling of GPCRs

4.11.1. Targeting the Naturally Occurring Functionalities in GPCRs. As all proteins utilize the same set of amino acids as the basic building blocks, strategies for labeling one of the 20 natural amino acids cannot afford a good selectivity in the cellular context crowded with biomolecules possessing similar reactivities. Targeting natural amino acids is practically limited to in vitro experiments. Covalent labeling of natural amino acid residues has been extensively studied and reviewed elsewhere.174,348 These methods rely on conjugation chemT

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distances between the seven transmembrane helices and the additional cytoplasmic helix were determined with highresolution and measured from 16 pairs of double spin-labeled rhodopsin mutants.367 This helix movement model was later quantitatively confirmed by the crystallographic studies on rhodopsin.120 Further comparison with the active state structures for β2AR,118 the adenosine A2A receptor,123,124 the M2 muscarinic acetylcholine receptor,126 and μOR128 elucidated a conserved molecular mechanism for GPCR activation. Along with the triumph of EPR spectroscopy, fluorescence techniques have been making strides in the GPCR field. In 1972, FRET experiment with rhodopsin performed by the Stryer group368 was the first study to shed light on the physical dimension of a GPCR. The fluorescent labeling was carried out without knowing the specific location of the modified cysteines, as the primary structure of rhodopsin would not be disclosed until 1983.369 It is worth noting that this study was among the earliest demonstrations of FRET as a molecular ruler for biological macromolecules.370 Since the 1990s, fluorescence techniques have been widely used to understand the structure− function relationship in rhodopsin. The native ligand 11-cisretinal can quench the fluorescence signal of the intrinsic tryptophan fluorescence.371 This quantitative assay for measuring the ligand binding and unbinding kinetics has been applied to probe the ligand binding pathway in rhodopsin.372−377 However, the usefulness of tryptophan is inherently limited by its near-UV wavelength and relatively low quantum yield. Cysteine labeling chemistries enabled a variety of environment-sensitive fluorescent probes to be site-specifically attached to rhodopsin. These studies showed that TM6 played a pivotal role in receptor activation by creating a hydrophobic crevice for engaging the C-terminus of G protein.378−381 Time-resolved transient fluorescence spectroscopy elucidated the sequence of events in the course of rhodopsin activation.382 Certain extrinsic fluorophores can be combined with the intrinsic tryptophan or tyrosine to make energy transfer pairs that offer better spatial resolution than the typical FRET scheme involving two extrinsic fluorophores.383,384 This fluorescencebased distance mapping method has been applied to assess the conformational heterogeneity of rhodopsin in different reconstitution systems.385 Cysteine labeling of β2AR with environment-sensitive fluorescent probes revealed a helix-movement activation mechanism, which was consistent with the earlier observations from rhodopsin.100,101 Fluorescence experiments showed that binding of partial agonists and full agonists with β2AR results in distinct conformational changes.102 The complete activation of the receptor involved multiple kinetic steps,103 and disruption of more than one conformational switch.104,386 Single-molecule studies on β2AR further revealed the dynamics of interconversion between different conformational states (see section 5.3).27,387,388 NMR is another powerful type of spectroscopy for elucidating protein structure and dynamics. There are two strategies for labeling the POI with NMR-active isotopes: metabolic incorporation of isotope-labeled amino acids, and covalent attachment of isotope labels to reactive residues. β2AR metabolically labeled with 13C-methionine was used to examine the activation dynamics.193,389 A particularly interesting isotope for protein NMR is 19F. Its large gyromagnetic ratio and 100% natural abundance make 19F NMR a highly sensitive NMR method (83% sensitivity relative to proton). 19F probes have

istries targeting the reactive groups of the amino acid residues, such as the primary ε-amine of lysine, the thiol group of cysteine, the side chains of arginine, histidine, tyrosine, and tryptophan, the N-terminal amino group, and the C-terminal carboxylic group. However, except for the N-terminal amino group and the C-terminal carboxylic group, none of these reactive residues is likely to occur only once in the protein. While steric factors or pH can differentially modulate the reactivities of different residues of the same type in one protein, and some selectivity may be achieved by a judicious choice of reaction conditions, the optimization process can be timeconsuming. The chemistries targeting sulfhydryl groups, such as disulfide exchange, alkylation using an alkyl halide (e.g., iodoacetamides), and Michael addition with α,β-unsaturated carbonyl compounds (e.g., maleimides), have been a popular choice to label GPCRs (Figure 17a). Cysteine is one of the least frequent amino acids in the composition of proteins. On the basis of the crystal structures of 32 unique GPCRs, 51% of cysteines are located in the transmembrane region, 34% are in the extracellular region, and the remaining 15% are intracellular.349 The majority of the extracellular cysteines form disulfides, which, together with the transmembrane cysteines, are resistant to the hydrophilic thiol-reactive reagents. The intracellular cysteines in the C-terminal tail may carry post-translational modifications (S-palmitoylation),350 further reducing the number of reactive cysteines in the receptor. Moreover, reactions of thiols with reagents, such as maleimides and iodoacetamides, require formation of a thiolate, which is more difficult in the hydrophobic transmembrane environment. Therefore, the labeling chemistries targeting cysteine thiols are particularly suited for labeling the cytoplasmic surface of GPCRs. Not surprisingly, tremendous efforts have been made to understand cysteine chemistry to create some minimalcysteine constructs for the prototypical GPCRs, rhodopsin351−356 and β2AR100,234,357,358 (Figure 17b). Lysines occur more frequently than cysteines in proteins, especially on the solvent-accessible surface. Consequently, labeling chemistries targeting lysine are not as selective, and less utilized than those targeting cysteines. Similar to cysteine labeling, lysine labeling involves the generation and validation of mutants. A particular concern for lysine mutants is that whether the substitution would alter its charge state and even disturb protein folding. 4.11.2. Spectroscopic Studies on GPCRs Enabled by Cysteine and Lysine Labeling. Biophysical probes have been attached to native or engineered cysteines to enable spectroscopic studies on GPCRs (Figure 17c). Site-directed spin labeling through cysteine chemistries has greatly facilitated the EPR study of GPCRs, in particular of rhodopsin. EPR spectroscopy was initially used to probe the protein−lipid interaction between rhodopsin and the rod outer-segment membranes.359,360 Its most important application, however, has been in distance mapping of receptor conformational changes. Such experiments required a single or a pair of nitroxide labels to be site-specifically attached to the receptor through cysteine chemistry. This approach enabled extensive spin-labeling studies of the cytoplasmic surface of rhodopsin,361−366 and provided the key insight that rhodopsin activation requires rigid-body helix movement characterized by the outward tilt of TM6.98 A remarkable application of doubly labeled GPCRs is the double electron−electron resonance (DEER) spectroscopy study of rhodopsin activation. In this study, the relative U

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Figure 18. Incorporating uaas into GPCRs by amber codon suppression. Functional GPCRs are typically expressed in eukaryotic cells. The uaacharged suppressor tRNA can be provided by either of the following two approaches: (1) the suppressor tRNAs are chemically acylated in vitro, and then delivered to the cells by microinjection or microelectroporation;416,665 (2) the cells are cotransfected with constructs encoding the evolved tRNA/aaRS pair and cultured in the presence of the cognate uaa. The bacterial tRNA/aaRS pair is orthogonal to the endogenous eukaryotic tRNAs and synthetases. The ribosome is capable of utilizing the suppressor tRNA charged with the uaas resulting in a full-length receptor tagged with a uaa at the desired position. Figure adapted and reprinted with permission from ref 666. Copyright 2011 Elsevier. Uaas 1−5 were incorporated into GPCRs using the chemical acylated tRNA approach,417,499 and uaas 6−9 by the orthogonal tRNA/aaRS pair.432,476,484

extensively used in studying ion channels and to a lesser extent in GPCRs.396,397 In most experiments, the extent of cysteine labeling was assessed by its absorption spectrum or fluorescence signal. The cysteine-reactive isotope label deuterated N-ethyl-maleimide enabled mass spectroscopy to be applied to measure cysteine accessibility.398,399 Lysine labeling, while less popular, is useful for probing receptor conformational change when the detection scheme, for example, NMR, can resolve the signals from individual lysines. In a 13C NMR study, the lysines of β2AR were labeled with 13C through reductive methylation. This modification preserved the positive charge of a Lys-Asp salt bridge linking the second and the third extracellular loops of β2AR, and thus made it possible to track the ligand-induced conformational change on the extracellular surface.400 Mass spectroscopy of β2AR labeled at the lysine side chains has also been reported.399 Altogether, the spectroscopic experiments invalidated the simplistic view that receptor activation, similarly to a toggle switch, merely involves two “on” and “off” states. A more accurate description treats GPCRs as a heterogeneous population comprised of energetically different conformations,

been chemically attached to the engineered cysteines in rhodopsin390−392 and in β2AR.393,394 The Overhauser effect of 19F pairs was exploited in mapping the conformational constraints in rhodopsin.391 19F NMR studies on β2AR provided evidence that ligands modulated the relative distribution of the heterogeneous conformational states and that both extracellular and intracellular stabilization were required for full activation of the receptor.393,394 Recently, 19F NMR experiments on the adenosine A2A receptor revealed a similar conformational selection mechanism.395 However, the labeling strategy determined which region of the receptor could be probed. The attachment of cysteine-reactive 19F probes was restricted to the cytoplasmic surface of the receptor. By comparison, metabolically incorporated 13C-methionine could be targeted to the transmembrane region. For both 13C NMR and 19F NMR experiments, the minimal methionine or minimal cysteine constructs were essential to simplify the spectrum. Differences in cysteine accessibility reflect the change of conformation in GPCRs, a fact that has been learned through the pioneering study on rhodopsin by George Wald.351 The substituted-cysteine accessibility method (SCAM) has been V

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most useful in generating soluble globular proteins, such as βlactamase,404 T4 lysozyme,412 staphylococcal nuclease,413 cytochrome P450 (CAM),414 etc. However, synthesis of functional membrane proteins requires the coordinated actions of multiple organelles to fold the polypeptides, add post-translational modifications, and transport them to the correct membranes, which can be very difficult to reconstitute in vitro. Because of the technical difficulty of delivering chemically acylated suppressor tRNA into cells, initially in vivo suppression was possible only for Xenopus oocytes whose large size facilitates injecting the tRNAs.415 Later the chemically acylated tRNAs were introduced into mammalian cells by microelectroporation (Figure 18: 1−5).416 These technical improvements enabled the Dougherty group to carry out an elegant study on the cation−π interaction involved in ligand binding to the M2 muscarinic acetylcholine receptor, and the D2 dopamine receptor expressed in Xenopus oocytes using progressively fluorinated unnatural amino acids.417 This approach can, in principle, be applied to a variety of GPCRs utilizing biogenic amine ligands. However, manipulating Xenopus oocytes and performing microelectroporation on mammalian cells require instruments and expertise unavailable in most laboratories. The Schultz group sought to develop a method for aminoacylating the suppressor tRNA in living cells. Their answer was to create an orthogonal suppressor tRNA/ aminoacyl-tRNA synthetase (aaRS) pair. In a seminal study published in 1981, Martin et al. showed that coinjection of yeast mitochondrial tRNA and bacterial aaRS into Xenopus oocytes led to the readthrough of the opal codon through the insertion of a tryptophan.418 This discovery implied that the suppressor tRNA/aaRS pairs could function in the heterologous host cells. In human complete protein coding genes, the amber codon is the least frequently utilized one among the three stop codons (amber, 23.5%; ochre, 29.4%, opal, 47.1%).419 In mammalian cells, amber codon suppression was found to produce readthrough proteins than opal and ochre suppression, partially because the aaRS aminoacylates the amber-suppressor tRNA more efficiently.4 These facts suggest that the amber codon is better suited than the other two for coding the 21st amino acid in cells. The engineering challenge was to establish a screening system to identify such an orthogonal pair that would utilize uaas and read the amber codon.420 The amber suppressor tRNA should not be the substrate for any endogenous aaRS, and yet be compatible with the endogenous ribosome. Also, the aaRS should specifically acylate the suppressor tRNA but not any endogenous tRNAs. Finally, cells should efficiently take up the uaas. In 2001, Wang et al. reported a Methanococcus jannaschii tyrosyl-tRNA/aaRS pair that fulfilled all of the above criteria to site-specifically incorporate phenylalanine analogues into proteins expressed in E. coli.421−423 Both the evolved suppressor tRNA and the aaRS were expressed inside the host cell using expression vector. In 2002, a new pair of mutant E. coli tyrosyl-aaRS and B. stearothermophilus suppressor tyrosyltRNA was generated to enable amber suppression in mammalian CHO cells.424 Further engineering resulted in site-specific incorporation of phenylalanine analogues into proteins expressed in yeast,425−427 Xenopus oocytes,428,429 insect cells,430 and various mammalian cell lines, including CHO cells,424,431 HEK293T cells,431,432 neurons,433 as well as neuronal stem cells.434

whose equilibrium is specifically dependent on the binding of unique ligands and G protein.135,136 This conceptual framework is significant not only for understanding the structure−function relationship of GPCRs, but also for the rational design of drugs. 4.11.3. Limitations to Targeting Naturally Occurring Functionalities. Although cysteine and lysine labeling has made a substantial contribution to understanding GPCRs, this approach is not without limitations. Reactive amino acids exist in abundance in cells, thereby limiting the feasibility of cysteine and lysine chemistries practically to purified receptors. However, as GPCRs possess multiple cysteines and lysines that participate in maintaining the seven-helix scaffold, ligand binding, and receptor activation,401 generating the minimal cysteine or lysine background constructs and validating the functional integrity of the mutant receptors remains a necessary and cumbersome process. There is no completely reliable method for predicting the reactivities of either cysteines or lysines. The labeling chemistries normally involve hydrophilic reagents, while any GPCR invariably contains a hydrophobic transmembrane helix bundle that is shielded by detergent micelles or lipid membranes. Most lysines reside in the exposed extracellular or intracellular surface, which makes them generally accessible by labeling reagents. Cysteines in the transmembrane region are often less amenable to modification. Nonetheless, labeling of transmembrane cysteines has been reported under certain conditions.100 Also, as palmitoylation of cysteine undergoes dynamic turnover,402 the palmitoylated cysteines in GPCRs are not absolutely resistant to cysteine chemistry. The long histories of studies on rhodopsin and β2AR have produced invaluable experiences for manipulating these two prototypical GPCRs and characterizing the functionality of their mutants, a legacy not readily available to researchers working on an expanding list of therapeutically interesting receptors. The difficulty with labeling the transmembrane regions of GPCRs also imposes a constraint on the application of spectroscopic methods. 4.12. Novel Approaches for Site-Specific Labeling of GPCRs

4.12.1. Incorporating Unnatural Amino Acids into GPCRs. The strategy to overcome the limitations of cysteine labeling in GPCRs was to employ genetically encoded unnatural amino acids (uaas).206,403 Among the 64 genetic codons, 61 code for amino acids, while the remaining three, opal (UGA), ochre (UUA), and amber (UAG), trigger termination of translation. In 1989, the Schultz group first successfully incorporated uaas into proteins by reassigning the amber codon to code for phenylalanine analogues.404 Their approach, known as amber codon suppression, was inspired by the earlier discoveries of amber suppressors in E. coli405 and yeast.406,407 Amber suppressors are certain tRNA species that are capable of suppressing the translation termination signal of the amber codon to yield a readthrough polypeptide. In eukaryotic cells, the complex of two polypeptide chain release factors, eRF1 and eRF3, mediates translation termination.408 The release factor complex specifically recognizes the stop codons to promote ribosome-catalyzed peptidyl-tRNA hydrolysis.409 The suppressor tRNA functions by competing against the release factor eRF1/eRF3 complex for binding with the stop codons in the mRNA transcript. The Schultz group developed efficient protocols for chemically aminoacylating the suppressor tRNA with uaas. The uaa-charged suppressor tRNAs could be utilized by the ribosome in a reconstituted translation system.404,410,411 This in vitro methodology was W

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In addition to the tyrosyl-tRNA/aaRS pairs, leucyl-,435 glutaminyl-,4,436,437 and typtophanyl-438 tRNA/aaRS pairs have also been reported. Nonetheless, the most facile system turned out to be the pyrrolysyl-tRNA/aaRS pairs.439−443 Selenocysteine (Sel) and pyrrolysine (Pyl) are known as the natural 21st and 22nd amino acids in proteins. Selenocysteine is coded by the opal (UGA) codon, and pyrrolysine by the amber (UAG) codon in certain organisms. Pyrrolysine is utilized in ribosomal protein synthesis in Methanosarcinaceae.444,445 The pyrrolysyl-aaRS displays remarkable side-chain promiscuity for the amino acid substrates, thus greatly expanding the chemical space of genetically encodable functionalities.446 Therefore, the directed evolution of pyrrolysyl-aaRS can be performed in E. coli and easily transferred to other systems. Structural studies showed that the contact surface between the pyrrolysyl-tRNA and the cognate aaRS is distinct from all of the other known tRNA/aaRS pairs,447 which underlies the excellent orthogonality of pyrrolysyl-based system in all of the bacterial and eukaryotic host cells tested.443 A comprehensive list of the pyrrolysine-derived uaas has been reviewed previously.448 The discovery and engineering of the pyrrolysyl system paved the way for uaa incorporation into multicellular organisms, including Caenorhabditis elegans,449 Drosophila melanogaster,450 and Arabidopsis thaliana.451 Recently, the pyrrolysyl-tRNA/aaRS pair was integrated into the genome of HEK293 cells, embryonic fibroblasts, and embryonic stem cells.452 These stable cell lines enable pyrrolysine analogues to be incorporated with >50% efficiency. The remarkable progress in cellular incorporation of uaas has been extensively reviewed.443,448,453−457 Despite the success of incorporating single uaas into proteins, progress in methods to simultaneously introduce two or more uaas has been slower. A major hurdle for simultaneous incorporation of multiple uaas is the suppression efficiency. The kinetic competition between stop codon suppression by the tRNA/aaRS pairs and normal chain termination by stop codon action constrains the incorporation efficiencies to 10− 20%. Consequently, the chance of obtaining a protein carrying two uaas from stop codon suppression is only a few percent.458 To solve this problem, a mutant version of the polypeptide release factor eRF1 was made that competes less effectively with the amber codon suppressor but modestly increased readthrough of opal and ochre codons. This engineered eRF1 was combined with a pyrrolysyl tRNA/aaRS pair to enable efficient incorporation of one uaa into multiple sites of proteins recombinantly expressed in HEK293T cells.459 To incorporate two different uaas into distinct sites of one protein, a unique codon that is orthogonal to the amber codon must be assigned to code for the second uaa. Several strategies have been devised to overcome this difficulty. First, an orthogonal ribosome can be evolved to decode the amber codon as well as a series of quadruplet codons.460 Second, pyrrolysyl-tRNA can be repurposed to recognize the opal (UGA) and ochre (UUA) codons and combined with the tyrosyl-tRNA/aaRS pairs.461,462 Third, pyrrolysyl-tRNAs can be evolved to decode quadruplet codons.463 So far, all of the doubly uaa-tagged proteins have been expressed in E. coli. A foreseeable application of the doubly tagged proteins is to attach two fluorophores at defined sites to enable FRET studies of protein conformational changes.464 Most GPCRs need to be expressed in eukaryotic cells, like mammalian or insect cell lines. Whereas functional expression of GPCRs in E. coli has been reported,465−467 the challenge for

refolding the receptor is nontrivial. The choice of expression system is further limited by the cell-based activity assays for the GPCR of interest. Therefore, those uaas compatible with eukaryotic expression systems are more serviceable to the GPCR field. In 2008, the first examples of incorporating uaas into GPCRs have been reported for the pheromone receptor Ste2p in yeast468 and for rhodopsin and the CC chemokine receptor 5 (CCR5) in mammalian cells.432 The efficiency of amber codon suppression in mammalian cells was improved by creating a novel chimera of H. sapiens and B. stearothermophilus tyrosyltRNA that forms an orthogonal pair with the existing E. coli Tyr-aaRS in the human HEK293T cell line.206,432 Uaas can be specifically inserted into the intracellular, extracellular, and transmembrane region of GPCRs, as long as the original residues are not structurally or functionally critical. Apart from rhodopsin, uaas have been incorporated into 35 discrete sites in the chemokine receptor CCR5,469−471 35 sites in the corticotropin-releasing factor receptor type 1,472 and 34 sites in the neurokinin-1 receptor (NK1R),473 all expressed in HEK293T or HEK293F cell lines. An exception was the ghrelin receptor, for which uaa-tagged mutants have been successfully expressed in E. coli and subsequently refolded.474 4.12.2. Genetically Encoded Unnatural Amino Acids as Biophysical Probes for GPCRs. There are two unique advantages when using genetically encoded uaas as spectroscopic probes. First, uaas can be incorporated into the TM core of GPCRs, which is not accessible by most labeling chemistries. Second, they do not require a reactive moiety for anchoring the probe, thus affording a shorter linker length between the probe and the protein backbone. Cornish et al. first reported site-specific incorporation of biophysical probes into in vitro synthesized T4 lysozyme.475 Among the uaas that have been incorporated into GPCRs, the azido group in p-azido-L-phenylalanine (azF, 6) serves as an excellent IR probe. Its vibrational signature between 2100 and 2150 cm−1 reflects the local electrostatic environment. This IR peak is distinct from that of the natural functionalities in proteins. The engineered azido-tagged rhodopsin has been used in Fourier transformation infrared (FTIR) difference spectroscopy experiments to track the conformational changes in the course of rhodopsin activation.476,477 Uaas can be used as cross-linkers for mapping ligand− receptor interactions, as well. azF (6) and p-benzoyl-Lphenylalanine (BzF, 8) are both photoactivatable cross-linkers and have been used to map the ligand binding modes for several GPCRs, including the chemokine receptors CCR5 and CXCR4,469,478−480 the corticotropin releasing factor receptor,481 and the neurokinin-1 receptor.473 A similar strategy was used to map the binding sites of conformation-sensitive monoclonal antibodies on the chemokine receptors CCR5 and CXCR4.482 A crystallographic study of the BzF photocross-linking product showed that the cross-linking is highly specific for proximal aliphatic chains.483 The application of targeted photo-cross-linking in membrane proteins has been reviewed previously.403,480 In many cases, it is impractical, though, to identify the crosslinking target of azF and BzF by crystallography and even mass spectrometry. The Wang group developed the thiol-reactive uaa p-fluoroacetyl-L-phenylalanine (Ffact, 9) that selectively reacts with a known proximal cysteine.484 The proximity-dependent cross-linking between Ffact and a cysteine-containing ligand yielded distance constraints for a 3D model of the ligandX

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between the strong infrared absorbances of the native functionalities. However, the nitrile probe exhibits a weaker signal (25−50%) than the azido probe.489,490 Genetically encoded uaas may also greatly facilitate the NMR and EPR experiments with GPCRs. L-4-Trifluoromethylphenylalanine (Figure 19: 14)491,492 results in a shorter distance between the NMR-active group and the protein backbone than 13 C-methionine and 19F-BTFA-labeled cysteine.389,394 The nitroxide uaa (Figure 19: 15),493 as compared to PROXYLlabeled cysteine,98 increases the linker length by three covalent bonds. Nonetheless, a unique advantage of these two uaas is that they can be inserted into the ligand-binding pocket and transmembrane bundle while cysteine labeling is largely confined to the cytoplasmic surface of the receptor. Uaas that function as photoinduced electron transfer (PET) acceptor can serve as conformational probes. p-Nitro-Lphenylalanine (Figure 19: 16) was shown to quench tryptophan fluorescence in a distance-dependent manner.494 Recently, 4-fluoro-3-nitrophenylalanine (Figure 19: 17) was developed as an ultrafast (picoseconds) PET quencher for the chromophore of GFP.495 Azobenzene is a well-described photoswitchable group. At room temperature, the trans isomer is the predominant species, as it is more stable than the cis isomer by 10−12 kcal mol−1. Irradiation at 340 nm triggers the trans−cis isomerization, pulling the two para-carbons closer by 3.4 Å. The transconformation can be restored through thermal relaxation or irradiation at 420 nm. Azobenzene-containing uaas (Figure 19: 18−21) provided the possibility of manipulating protein activity by light.496−498 The photoswitchable uaas (Figure 19: 19−21) with reactive handles were developed for a mammalian expression system.498 These photoswitchable uaas can be exploited to modulate the accessibility of the ligand-binding pocket. For example, uaa 21 has been shown to form a covalent linkage with a proximal cysteine to constitute an additional structural constraint. This optogenetic approach may provide mechanistic insights into receptor conformational changes and enable the development of photoactivatable GPCRs. The smallest useful intrinsic fluorescent probe in proteins is tryptophan because the fluorescence of phenylalanine, histidine, and tyrosine is too weak for practical applications in biochemistry. However, its ubiquitous presence in proteins makes it impossible to analyze a particular protein in the cellular context by tryptophan fluorescence. Also, tryptophan suffers from a relatively low quantum yield (about 0.2), a relatively low extinction coefficient, and the need for damaging UVB excitation (280−315 nm). Hence, there has been continuing efforts in developing fluorescent uaas with redshifted spectra and higher brightness (the brightness being defined as the product of extinction coefficient and fluorescence quantum yield). Before genetic incorporation of uaas was made possible, several fluorescent uaas had been synthesized, chemically ligated to the suppressor tRNAs, and incorporated into proteins either expressed in Xenopus oocytes or synthesized in cell-free translation systems. 3-N-(7-Nitrobenz2-oxa-1,3-diazol-4-yl)-2,3-diaminopropionic acid (Figure 20: 5, NBD-Dap) was used as a FRET donor to detect ligand binding to the neurokinin-2 receptor (NK2R),499 which was also the first example of incorporating a fluorescent uaa into a membrane protein. The electrostatic reporter Aladan (Figure 20: 22) made it possible to probe the interior environment of potassium channels and of an IgG-binding domain.500 BODIPY phenylalanine analogues have been incorporated into strepta-

binding mode between the corticotropin-releasing factor receptor and its native peptide ligand urocortin I, in the absence of any crystal structure.472 Even in an era when the momentum of GPCR crystallography is vividly felt, crystallizing a GPCR remains a daunting task. More importantly, many native ligands simply do not have the high affinity to effectively stabilize the receptors. In fact, structures of GPCRs in complex with their native ligands have been described, but only for rhodopsin and β2AR.106,196 Thus, targeted cross-linking nicely complements crystallography in understanding receptor−ligand binding. In addition to the nine uaas described in Figure 18, several other uaas are potentially beneficial to research on GPCRs. Diazirine uaas with different linker length (Figure 19: 10, 11, 12) may enrich the targeted photo-cross-linking methodology.485−487 A nitrile-uaa488 can serve as an IR probe (Figure 19: 13). Similar to azF, the environment-sensitive nitrile vibrational signal (2200−2250 cm−1) falls into a spectrally clean window

Figure 19. Other genetically encoded biophysical probes. 10: Photocross-linking tyrosine analogues containing a diazirine group.485 11,12: Aliphatic diazirine uaas with different linker lengths.486,487 13: Cyanouaa as IR probe.488 14: 19F-uaa as NMR probe.491 15: Nitroxide uaa as EPR probe.493 16,17: Nitro-uaa as PET quencher.494,495 18: Photoswitchable azobenzene-uaa.496 19−21: Photoswitchable azobenzene-uaas with a terminal alkene, ketone, and chloroalkane reactive handle, respectively.498 Y

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4.12.3.1. Targeting Genetically Encoded Ketone Handles. The first example of site-specific bioorthogonal protein modification was achieved for T4 lysozyme containing a ketone uaa.514 To import this approach into the GPCR field, the wellexpressed rhodopsin was chosen as a model system to test the strategy of labeling genetically encoded bioorthogonal reactive handles. The binding of its inverse agonist 11-cis-retinal confers rhodopsin excellent thermal stability.515 An unexpected finding was that the wild-type rhodopsin exhibited a substantial level of nonspecific reactivity toward reagents targeting the ketone group.432,510,516 Thus, the ketone handle is not strictly bioorthogonal. A plausible explanation is that GPCRs may undergo unexpected carbonylation due to the cellular oxidative stress,517−519 and the level of carbonylation can be contingent on the expression system. As the ketone moiety is generally believed to qualify as a bioorthogonal reactive handle, this finding raised some concern in the bioconjugation field. 4.12.3.2. Targeting Genetically Encoded Azide Handles. 4.12.3.2.1. Copper-Catalyzed Azide−Alkyne [3+2] Cycloaddition (CuAAC). Unlike ketones and aldehydes, azides are entirely absent from living systems and stand out as more promising candidates for bioorthogonal labeling. A variety of chemistries targeting azides have been described.520−522 Among them the most notable example is copper-catalyzed azide− alkyne [3+2] cycloaddition (CuAAC), also widely known as copper-catalyzed “click” chemistry.523,524 The concept of “click chemistry” was popularized by Sharpless et al. in 2001 to describe an ideal type of reactions that is modular, wide in scope, stereospecific, generates only inoffensive byproducts that can be removed by nonchromatographic methods, and proceeds under simple and mild conditions.525 CuAAC satisfies most of these criteria and was soon applied to bioconjugation.526 However, the presence of transition metals may disrupt the native function of proteins by coordinating with cysteine residues, or by creating free radicals that lead to protein backbone cleavage or side chain damage, let alone the cellular toxicity.527 Whereas it is possible to reduce the undesirable consequences by using metal-chelating ligands528 or shortening the reaction time, it is debatable whether these strategies would be cost-effective for labeling GPCRs. In our initial attempt to label azF-rhodopsin using CuAAC, we observed some backbone cleavage (unpublished data). Therefore, this Review will only describe metal-free bioorthogonal chemistries. Similarly, this Review will not cover the click chemistries catalyzed by palladium for bioconjugation of purified proteins or in cells.529,530 4.12.3.2.2. Staudinger−Bertozzi Ligation. Staudinger− Bertozzi ligation involving modified phosphines (IUPAC: phosphanes) was the first bioorthogonal chemistry for azides.531 Fluorescent labeling of azF-tagged rhodopsin and CCR5 has been demonstrated.470,516 The Staudinger ligation gave clean background, but the reagent was susceptible to oxidation, and the reaction was too slow (k2 = 10−3−10−2 M−1 s−1) to give stoichiometrically labeled receptor. Moreover, a significant level of noncovalent binding between the receptor/ micelle and the phosphine makes it difficult to remove the excess labeling reagents under the mild conditions for maintaining receptor functionality.516 4.12.3.2.3. Strain-Promoted Azide−Alkyne [3+2] Cycloaddition (SpAAC). The spontaneous reactivity of strained cyclooctynes with azides was initially described in the 1950s and 1960s.532−534 The Bertozzi group first recognized the potential of strain-promoted azide−alkyne [3+2] cycloaddition

Figure 20. Fluorescent amino acids. Upper row: Tryptophan and uaas chemically loaded onto the suppressor tRNAs and incorporated into membrane proteins expressed in Xenopus oocytes. 5: NBD-Dap was the first fluorescent uaa encoded into a GPCR.499 22: The electrostatic reporting uaa Aladan was incorporated into potassium channels and IgG-binding domain.500 23: (BODIPYFL)K has been incorporated into the nicotinic receptor.503 Lower row: Genetically encoded fluorescent unnatural amino acids. 24: 5-OH-Trp.504 25: HceG containing hydroxycoumarin.505 26: DansA containing dansyl alanine.506 27: Anap containing acetyl naphthalene.508 Only 24 and 27 have been shown for amber codon suppression in mammalian cells.

vidin and calmodulin using a quadruplet codon in a cell-free translation system.501,502 A BODIPY lysine analogue (Figure 20: 23, (BODIPYFL)K) enabled single-molecule fluorescence detection of the uaa-tagged nicotinic receptor on oocytes.503 Because these fluorescent uaas need to be chemically charged to the suppressor tRNA, they are more useful for membrane proteins that can be studied in Xenopus oocytes. So far four genetically encoded fluorescent uaas have been reported, including 5-hydroxyl-L-tryptophan (Figure 20: 24, 5OH-Trp),504 L-hydroxycoumarin ethylglycine (Figure 20: 25, HceG),505 dansylalanine (Figure 20: 26, DansA),506 and 3-(6acetylnaphthalen-2-ylamino)-2-aminopropanoic acid (Figure 20: 27, Anap).507,508 Among them, 5-OH-Trp and Anap have been successfully used in mammalian cells. 4.12.3. Bioorthogonal Labeling of GPCRs Targeting Genetically Encoded Reactive Handles. The property of the substrate-binding pocket of the aaRS imposes a practical constraint on the size and consequently on the photophysics of any fluorescent uaa that can be genetically encoded in cells. Therefore, there is a pressing need for developing bioorthogonal labeling strategies for attaching the larger, longerwavelength fluorophores, such as cyanine or rhodamine, to GPCRs. Our group has developed a two-step strategy to label GPCRs: reactive handles are first genetically encoded into the receptors using amber codon suppression, and then reacted with bioorthogonal chemistries.206,432,470,471,509−512 This approach is more generalizable than cysteine labeling, as it eliminates the need for creating a minimal-cysteine construct and ensures a high site-specificity. Because outstanding reviews on bioorthogonal chemistries are available,174,348,457,513 this work shall focus on their applications for labeling GPCRs. Z

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Figure 21. Bioorthogonal labeling of uaa-tagged GPCRs. (a) The uaa-tagged GPCRs heterologously expressed in a eukaryotic cell (i) can be labeled by bioorthogonal chemistries on cells (ii). The nonspecific action of “leaky” tRNA/aaRS pairs may result in a low level of full-length receptor without the bioorthogonal reactive handle (iii). When the amber codon is positioned close to the N-terminus, internal translational reinitiation may result in a folded receptor with an incomplete N-terminus (iv). The proteasome can degrade the truncated peptides terminated at the amber codon (v). (b) Bioorthogonal labeling chemistries targeting the ketone and azide functionalities. p-Acetyl-L-phenylalanine (AcF, 6) reacts with hydrazone and oxime reagents. p-Acetyl-L-phenylalanine (azF, 7) reacts with phosphine (Staudinger−Bertozzi ligation), terminal alkynes (copper-catalyzed azide−alkyne [3+2] cycloaddition), or cyclooctynes (the strain-promoted azide−alkyne [3+2] cycloaddition, SpAAC). (c) Experimental scheme for labeling uaatagged GPCRs in vitro. The heterologously expressed receptor is solubilized in detergent and immunopurified with the C-terminal specific antibody. The receptor bound to the resin is subjected to the labeling reaction. In the end, the labeled receptor is specifically eluted from the resin. (d) The sites of uaa incorporation and labeling in rhodopsin (ice blue, uaa incorporation; red, fluorescent labeling with SpAAC).476,477,510,511 (e) The absorption spectra of rhodopsin labeled with different Alexa Fluor dyes (normalized on the basis of the concentration of functional rhodopsin).509,510 AA

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Figure 22. Genetically encoded bioorthogonal reactive pyrrolysine analogues. Pyl: Pyrrolysine. 28,29: Aliphatic alkyne (AlkK) and azide (AzK).542 30: Acrylamide (AcrK).544 31: Cyclopropene (CpK).545 32: 1,2-Aminothiol (ThiPK).546 33,34: Cyclooctynes (CoK1, CoK2).547 35: Bicyclononynes (BCNK).548,553 36,37: Norbornenes (NorK1 and NorK2).550,551 38: trans-Cyclooctene (TCOK).552,553

(SpAAC) for bioorthogonal labeling.535 This reaction, also referred to as copper-free click chemistry, was widely appreciated for its value in bioconjugation. Much effort has been dedicated to enhancing the reactivity, stability, and optimization of the synthetic routes.522 Tian et al. showed that SpAAC using dibenzocyclooctyne (DIBO)536 served as a robust method for labeling azF-tagged rhodopsin.510 The modular design of the labeling reagent allowed a variety of fluorophores and peptides to be sitespecifically attached to the intracellular, extracellular, and transmembrane region (Figure 21d,e). The labeled receptor is functional with respect to activation and ligand binding.509−511,537 Apart from rhodopsin, the combination of DIBO and azF-tagged GPCRs has been demonstrated for CCR5471 and the ghrelin receptor.474,512 It is worth noting that the azF-tagged ghrelin receptor was either expressed in mammalian cells and purified from the plasma membrane,512 or expressed in E. coli and refolded to its native state.474 One of the known issues of cyclooctyne reagents is the tradeoff between hydrophobicity, nonspecific background reactions, and SpAAC reactivity. Cyclooctynes have been reported to react with thiols through a thiol−yne reaction.538,539 The selectivity factor of DIBO for azF over cysteine was estimated to fall between 200:1 and 800:1,510 which is sufficient for a chemically defined system. Some other cyclooctynes, for example, bicyclo[6.1.0]nonyne (BCN), exhibit greater crossreactivity with cysteine thiols, which limits the use of SpAAC for protein labeling.539 BCN is much less hydrophobic than DIBO, which reduces partitioning into membranes and nonspecific protein binding. It was recently observed that inclusion of a low concentration of β-mercaptoethanol suppresses the cross-reactivity between BCN and thiols without significantly compromising the efficiency of SpAAC for in vitro protein labeling.540 4.12.3.2.4. Micelle-Enhanced SpAAC. GPCRs have heterogeneous surface hydrophobicity. The SpAAC involving cyclo-

octyne reagents preferably labels the hydrophobic TM region of rhodopsin. The reaction rates for azido groups situated on the transmembrane surface of rhodopsin are accelerated by up to 103-fold (k2 > 100 M−1 s−1)511 as compared to the sites on water-exposed surfaces, or the literature value (k2 = 0.1−1 M−1 s−1).521,536 This observed rate enhancement was attributed to the amphiphilic nature of the labeling reagent, which is comprised of a hydrophobic cyclooctyne and a hydrophilic dye, resulting in partitioning of the labeling reagent into the micelles. Specifically, the cyclooctyne partitions into the hydrocarbon core of the detergent micelle where it results in a high local reactant concentration. This micelle-enhanced SpAAC reaction, first observed from labeling GPCRs, was supported by a later report involving small molecules.541 The experiences with labeling the CCR5 receptor provided similar insights. The exposed extracellular and intracellular regions were better labeled with the Staudinger reaction.470 In contrast, a residue located deep in a hydrophobic binding pocket was readily modified by SpAAC.471 Taken together, these findings shed light on how the local environment on the protein surface can modulate the efficiencies of protein labeling. The capability of targeting the TM region of GPCRs complements the cysteine labeling chemistry that mostly targets the intracellular region. 4.12.3.2.5. Applications of Site-Specific Bioorthogonal Labeling of GPCRs. Bioorthogonal fluorescent labeling of azF-tagged GPCRs enables new FRET-based assays to investigate the GPCR signaling complex. Alexa Fluor 488labeled rhodopsin was exploited in a fluorescence-quenching assay for monitoring the formation of mature pigments.510,511 The ghrelin receptor labeled with Alexa Fluor 647 at the extracellular end of TM4 facilitated the study of ligand binding (Figure 12k).512 The ghrelin receptor tagged with Alexa Fluor 488 at the intracellular end of TM1 was used to study the assembly of receptor−G protein complex (Figure 12m).474 In AB

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Figure 23. Bioorthogonal labeling reactions with pyrrolysine analogues. (a) “Photoclick” chemistry between genetically encoded acrylamide (12, AcrK) and tetrazole.451 (b) “Photoclick” chemistry between genetically encoded cyclopropene (13, CpK) and tetrazole.545 (c) Cyanobenzothiazole condensation with genetically encoded 1,2-aminothiol (14, ThiPK).546 (d) Strain-promoted azide−alkyne [3+2] cycloaddition (SpAAC) between genetically encoded cyclooctynes (15, CoK1; 16, CoK2; 17, BCNK).547,548 (e) Strain-promoted inverse-electron-demand Diels−Alder cycloaddition (SPIEDAC) between genetically encoded cyclooctyne and tetrazine.553 (f) SPIEDAC between strained alkenes (18, NorK1; 19, NorK2; 20, TCOK) and tetrazine.550−553

4.12.3.4. Bioorthogonal Labeling of uaa-Tagged GPCRs on the Cell Surface. Fluorescent labeling of uaa-tagged GPCRs on the surface of live cells has not been demonstrated, but can be readily envisioned. The difficulty of achieving good contrast for on-cell labeling depends on several factors: first, the expression level of the uaa-tagged GPCRs; second, the nonspecific reactivity of the labeling reagent that results in a covalent bond with the native chemical functionalities in cells; third, the nonspecific, noncovalent binding between the labeling reagent and the cell surface. GPCRs are relatively low expressing on the mammalian cell surface (103−106 copies/cell). It should be kept in mind that bioorthogonal chemistries have high, but not absolute, selectivity over the native functionalities. In fact, strained alkenes554 and strained alkynes539 were shown to react with thiols. The cross-reactivity between tetrazines and thiols has also been suggested.555 On the cell surface the abundance of cysteines (>108 copies/cell) is orders of magnitude higher than that of even a high-expressing GPCR.510 For example, in the case of SpAAC between azF and dibenzocyclooctyne, its selectivity factor for azF over cysteine is between 200:1 and 800:1. Even if only 1/10 of the cysteines in membrane proteins were available for modification by cyclooctyne, the resulting signal-to-noise ratio would be far from ideal. The hydrophobic binding between cyclooctyne and the lipid bilayer of the plasma

all of these examples, the freedom of choosing the labeling site allowed rational design of the energy transfer scheme. 4.12.3.3. Targeting Genetically Encoded Pyrrolysine Analogues. The rapid progress in the pyrrolysyl-tRNA/aaRS system has enriched the toolkit of genetically encodable bioorthogonal reactive handles (Figure 22, Figure 23). Successful examples include (but not limited to): aliphatic alkynes (AlkK, 28),542 aliphatic terminal alkynes (azK, 29),542,543 acrylamide (AcrK, 30),451,544 cyclopropene (CpK, 31),545 1,2-aminothiols (ThiPK, 32),546 strained alkynes (CoK1, 33; CoK2, 34; BCNK, 35),547−549 strained alkenes (NorK1, 36; NorK2, 37; TCOK, 38),550−552 etc. Some of these newly developed labeling methods exhibit ultrafast reaction kinetics, particularly the strain-promoted inverse-electrondemand Diels−Alder cycloaddition (SPIEDAC) between tetrazine and BCNK (Figure 23e; k2 = 103−104 M−1 s−1), or between tetrazine and TCOK (Figure 23f; k2 > 104 M−1 s−1).513,552,553 While these fast labeling reactions await experimental demonstration for GPCRs, they should be particularly useful for labeling receptors with a short lifetime. A potential issue is that some genetically encoded reactive handles like strained alkynes or alkenes may suffer from crossreactivity with proximal cysteines. Hence, the labeling site may require optimization. AC

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Figure 24. Fluorogenic labeling strategies. (a) Fluorogenic reaction scheme 1: the dye bears a quenching reactive group and becomes highly fluorescent when the reaction alters the quenching group. (b) Azide as the quencher. The fluorescence signal is turned on by reaction with terminal or strained alkynes. (c) Tetrazine as the quencher. The dye is turned on by reaction with strained alkynes or alkenes. (d) Examples of fluorogenic dyes: hydroxycoumarin-azide,561 HELIOS-400Me tetrazine,667 Oregon Green-tetrazine,564 BODIPY-tetrazine,568 fluorescein derivative-azide,566 Sirhodamine derivative-azide.567 (e) Fluorogenic reaction scheme 2: the reaction causes the quencher to dissociate. (f) SNAP-tag for fluorogenic labeling. The fluorophore is attached to the benzyl group and the quencher to guanine. In BG-DY549-QSY7, QSY7 quenches the fluorescence of DY549 by 98%,569 corresponding to 50-fold signal enhancement. (g) TMP-tag for fluorogenic labeling. The first TMP-based fluorogenic label, TMPQ-Atto520, exhibits 20-fold signal enhancement upon covalent binding.311 AD

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reinitiation may result in a correctly folded product that only misses a fragment in the N-terminal domain but still covers the rest of the original sequence.469 These two scenarios may give rise to “ghost” receptors that escape fluorescent labeling but still signal. Therefore, it is essential to validate, with the most sensitive assay available, that the amount of untagged receptor is truly irrelevant to the biological question at hand.

membrane only exacerbates the problem. This estimation suggests that using cyclooctyne reagents to label and image low-abundance molecules on living cells is challenging. Another issue to bear in mind is that GPCRs, like other membrane proteins, do not stay indefinitely at the cell membrane. Their surface residence time ranges from minutes to hours.175 While endocytosis blockers or reduced temperature can be used to prolong the residence time, they may interfere with subsequent experiments. The fast labeling chemistry targeting genetically encoded strained alkenes or alkynes may represent a solution for on-cell labeling. The azide and tetrazine moieties are less hydrophobic than cycloocytnes, which helps to reduce nonspecific binding to the membrane. Furthermore, azide and tetrazine fluorescent labeling reagents can be fluorogenic, in other words, exhibiting dramatic signal enhancement upon conjugation. The strategies for developing fluorogenic labeling reagents will be discussed in greater details in section 4.13. Fluorogenic labeling of genetically encoded strained alkenes or alkynes has been demonstrated for heterologously expressed insulin receptor.556 4.12.4. Potential Issues with Amber Codon Suppression in Living Cells. The application of amber codon suppression in living cells warrants some discussion (cf., Figure 21a). First, protein production for ensemble spectroscopic experiments routinely demands a significant amount of sample. For example, the FTIR studies on azF-tagged rhodopsin have taken advantage of the fact that purified functional rhodopsin harboring uaas can be obtained at submilligram scale from mammalian cell culture.476,477 Other uaa-tagged GPCRs had to be analyzed by more sensitive assays like photo-cross-linking and fluorescence.206,403 As stated earlier, prokaryotic cells are generally unable to express correctly folded and post-translationally modified GPCRs. Thus, the major bottleneck was the lack of efficient eukaryotic expression systems for amber codon suppression, which has limited the applicability of a wide range of uaas to bacterially expressed proteins. The newly developed stable cell lines with a tRNA/aaRS pair for amber codon suppression452 may prove advantageous for this purpose. Second, 23.5% of the endogenous genes also use the amber stop codon.419 The consequences of off-target amber codon suppression remain understudied. The viability of uaa-tagged cells and animals suggests that amber codon suppression does not cause severe cytotoxicity. However, any interference with the cellular signaling network cannot be excluded. Third, amber codon suppression for an overexpressed POI is incomplete (typical efficiency for a single mutation: 5−20% using the tyrosyl tRNA/aaRS pairs) due to the competition of the eRF1/eRF3 complex, yielding truncated polypeptides. Although the truncated peptides are likely to be misfolded and then degraded by the proteasome, the efficiency of such cellular quality-control machinery and the cellular consequences are unclear. Last, the nonspecific substrate usage of the tRNA/aaRS pair may produce full-length protein even without exogenous uaas. In the majority of the published reports, the “leakiness” of a tRNA/aaRS pair is typically determined by Western blot, whose sensitivity depends on the affinity of the primary antibody and the sample processing procedure. However, the absence of nonspecifically expressed protein in the Western blot does not preclude any detectable activity in cell-based assays, particularly in highly sensitive assays (e.g., patch clamp or luciferase reporters). Also, when the amber codon is positioned close to the N-terminus, internal translation

4.13. Fluorogenic Labeling Reactions

Fluorogenic reactions can be particularly useful for achieving high contrast in the complex cellular environment.557−560 There are two popular strategies for designing fluorogenic probes. In the first strategy, the reactive moiety serves as the quencher for the linked dye. Conjugation with the cognate reactive handle on the POI destroys its quenching effect (Figure 24a). The first example for fluorogenic click reaction involved azido-hydroxycoumarin,561 followed by a variety of azido-based fluorogenic dyes.557 Later tetrazine was found to fulfill the fluorogenic criterion as well.562−564 The azide group functions as a PET quencher for coumarins,561 anthracene derivatives,565 xanthene derivatives,566,567 etc. The 1,3-dipolar cycloaddition of an azide and a terminal or strained alkyne converts the azide into a triazole ring, resulting in the loss of PET quenching (Figure 24b). Similarly, the quenching effect of tetrazine group is deactivated upon reacting with strained alkenes or alkynes (Figure 24c).552,562,568 In the second strategy, the quencher and the dye are connected by a cleavable linker (Figure 24e). Upon reaction, the concomitant release of the quencher unmasks the fluorescence emission. The modular nature of SNAP-tag (Figure 24f) and TMP-tag (Figure 24g) substrates was harnessed to design the second type of fluorogenic probes.311,569,570 The quencher is attached to the leaving moiety so that only the fluorophore ends up attached to the receptor. 4.14. Choosing the Right Labeling Method To Understand the Biochemistry and Cell Biology of GPCRs

4.14.1. Tracking GPCR Conformational Change. Various spectroscopic methods have been applied to track the conformational change in the course of GPCR activation. The probes, together with the linker to the protein, should be as small as possible to report the movement of the polypeptide backbone faithfully. On the other hand, a longer linker may facilitate probe reorientation and reduce orientational artifacts in FRET-based assays. The probes should also give strong signals so that receptor expression is less likely to be a hurdle. In the past, such probes were typically attached to the receptor through chemistries targeting cysteine thiols. The development of genetically encoded unnatural amino acids has the potential of overcoming this classic approach. The uaas can be incorporated into the TM region of GPCRs that is not accessible by most labeling chemistries. If the probe is only a few atoms in size and can be incorporated as part of the uaas, the linker length between the probe and the protein backbone can be dramatically reduced. As for the probes that cannot be genetically encoded, bioorthogonal labeling of uaa-tagged GPCRs may offer a general solution for site-specifically attaching them to GPCRs. As compared to chemistries targeting cysteine, bioorthogonal labeling targeting uaas benefits from its greater freedom in choosing the site of labeling. The fluorogenic bioorthogonal labeling strategy may ultimately enable single-molecule fluorescence studies of receptor conformational changes in the cell membrane. AE

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Another approach for probing conformational change is to utilize conformational-specific biomolecules, such as a ligand, an antibody, or a nanobody. Preparing such reagents is nontrivial, nonetheless. In the existing literature, the GFPnanobody biosensor198 represents an interesting case. Expressing the nanobody biosensor in the cellular milieu enabled one to overcome the barrier of the plasma membrane, which may open new possibilities for tracking GPCR conformational changes in organelles. 4.14.2. Trafficking and Internalization. Any method resulting in stable and bright labeling of a GPCR is theoretically useful in tracking the cellular localization of the receptor. The most popular strategies are based either on an epitope-specific antibody or on a fluorescent protein fusion. For example, a quantum dot-labeled high-affinity antibody was used to visualize the internalization and endosomal trafficking of epitope-tagged serotonin receptors.571 The question is whether such modification may alter the native behavior of the receptors. Antibodies, fluorescent proteins, and self-labeling proteins are all not much smaller in size than GPCRs (Figure 25), which may lead to interferences with receptor function. In fact, alteration of receptor mobility caused by fluorescent protein tagging has been reported.572 Ligand-directed labeling has also been successfully applied to monitor the internalization of membrane proteins.224 However, the prerequisite of this approach is that the ligand itself will not induce receptor internalization. By comparison, bioorthogonal chemistry targeting uaa-tagged GPCRs would produce minimal modification of receptors. It would be of general interest to evaluate whether this strategy is suitable for tracking the cellular localization of GPCRs. 4.14.3. Oligomerization. As described in section 3.5, much remains to be elucidated about GPCR oligomerization. Figure 26 summarizes the methods for detecting GPCR oligomerization within the scope of this Review. Most of them utilize FRET between fluorescent probes attached to monomeric receptors (Figure 26a−f). As compared to the methods based on fluorescent antibodies or protein tags, using fluorescent ligands for imaging GPCR oligomers (Figure 26b,c) has two important advantages. First, it is possible to image endogenously expressed GPCRs in native tissue because there is no need to overexpress a modified receptor expression construct. Second, fluorescent ligands are smaller than antibodies or protein tags. However, the use of fluorescent ligands, particularly in the case of fluorescent bivalent ligands, is limited by the availability of such reagents. Proximitydependent enzymatic labeling methods, like ID-PRIME or BioID, report protein interactions based on the physical proximity of binding partners (Figure 26g). While there is but one example in the published literature that applied IDPRIME to detect GPCR oligomerization,344 this strategy has a great potential for understanding GPCR signaling networks. Apart from fluorescence techniques, the degree of oligomerization can be analyzed by chemical cross-linking/mass spectrometry, raising the possibility of profiling GPCR oligomerization.

Figure 25. Comparing the sizes of GPCR, proteins tags, and fluorescent reporters. All of the molecules, as well as the quantum dot, are shown in scale. The crystal structures of (a) an immunoglobulin G (IgG, PDB: 1IGT),668 (b) a representative GPCR rhodopsin (Rho, PDB: 1U19),109 (d) GFP (PDB: 1GFL),669 (e) SNAP-/CLIP-tag (PDB: 3KZY), (g) Halo-tag (PDB: 4KAA), (h) TMP-tag (PDB: 1DR7), and (i) Renilla luciferase (RLuc, PDB: 2PSD)670 were prepared using VMD.671 The IgG molecule and its Fab and Fc regions illustrate the size of typical labels for immunofluorescence. The chromophore of GFP is highlighted in orange, and the active sites for the SNAP-/CLIP-tag (C145), Halo-tag (D106), and TMP-tag (L28C) are in red. The molecular model for Alexa647 (c) was generated using Schrödinger Maestro. The transmission electron micrograph (f) shows the structure of a 12 nm (CdSe)ZnS core−shell quantum dot with far red emission, similar to Alexa647. Electron micrograph courtesy of Andrew M. Smith. Reproduced with permission from ref 672. Copyright 2010 American Chemical Society.

be labeled with a fluorescent dye precisely at a 1:1 molar ratio (and not just an average dye/protein ratio of 1:1) using the methods described in the previous section. Despite a long list of papers in which over 45 different GPCRs have been investigated, the majority of studies were focused on tracking GPCR diffusion in the cell membrane, largely due to the longstanding limitation in site-specific labeling of the receptors. With the recent introduction of site-specific labels via SNAP-tag technology or unnatural amino acids (Tables 3 and 4), GPCRs can be selectively labeled in the cellular complexity. Consequently, important questions related to GPCR biology, such as conformational dynamics, ligand-binding dynamics, or mesoscale organization of GPCRs in the cell membrane, can be tackled in a meaningful way.

5. APPLICATION OF SINGLE-MOLECULE METHODS TO GPCRs Single-molecule imaging is ideally suited to decipher the dynamic and heterogeneous behaviors of GPCRs. In this section, we review the extensive literature reports in which single GPCRs were observed (Table 1). To conduct singlemolecule fluorescence imaging of GPCRs, each receptor has to AF

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Figure 26. Detecting GPCR oligomerization in living cells. Most detection schemes are based on energy transfer between (a) fluorescently labeled antibodies, (b) two fluorescent ligands, (c) a fluorescent bivalent ligand, (d) fluorescent proteins, (e) luciferase and a fluorescent protein, and (f) fluorophores conjugated to the orthogonal SNAP-tag and CLIP-tag. GPCR oligomerization can also be probed by proximity-dependent labeling, for example, (g) Interaction-Dependent Probe Incorporation Mediated by Enzymes (ID-PRIME).

5.1. Mobility, Oligomerization, and Stoichiometry

of GPCRs does not seem to be a general requirement for ligand recognition or signaling, but it is speculated as a mechanism for the cell to modulate receptor mobility at the cell surface, receptor intracellular trafficking, or receptor signaling functions. Current models describe the plasma membrane as a complex dynamic heterogeneous distribution of lipids and proteins in which signaling from cell surface receptors is often highly compartmentalized, with receptors existing in signaling microdomains such as caveolae or lipid rafts whose organization is mediated by specific protein−protein or protein−lipid interactions. In this context, the lateral mobility of receptors is a key parameter describing how they might move in and out of such microdomains and encounter other identical or different receptors to form transient or stable dimers or oligomers. The lateral mobility of GPCRs was initially investigated by fluorescence recovery after photobleaching (FRAP), using receptors tagged with a fluorescent protein or labeled with a fluorescent ligand. In a FRAP experiment, a defined area of the

Many membrane receptors function as homo- or heterodimers, or even as higher order oligomers, and oligomerization confers unique properties that monomers lack. For example, each monomer may contribute to the formation of the ligand binding site or recruiting intracellular adapter proteins. For a long time it was thought that GPCRs were an exception among membrane receptors by functioning as a monomer. Singlemolecule colocalization imaging and step-photobleaching analysis provided unequivocal evidence that single monomeric β2AR and μOR molecules incorporated into membrane-mimic high-density lipoprotein particles were capable of binding and activating their G protein.144,573 Monomeric rhodopsin in solution was shown to activate its G protein transducin as well.147 Nonetheless, evidence has accumulated in parallel that GPCRs are also capable of forming dimers, although it is still far from clear when and where this process takes place under physiological conditions.151,574 Dimerization or oligomerization AG

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technique

AH

anti-FLAG-antimouse-F (ab′)2-QD anti-FLAG-antimouse-F (ab′)2-QD agonist-bodipy630 antagonist-bodipy630, A3AR-GFP α1AR-CFP, Gaq-citrine

SPT

SPT

mobility

SPT fs-XR AFM

AFM AFM

AFM

AFM AFM

stability stability

stability

stability stability

α2AAR AT1R bR

β2AR

β1AR

SPT

mobility

SMLM

secondary antibody-Cy2 or Cy3 secondary antibodyAlexa Fluor 488 β2AR-mEos2 fusion

NSOM

distribution

SNAP-Alexa Fluor 647

SPT

B2R-GFP

mobility, signaling mobility, oligomers oligomers

FCS

FCS

SPT

mobility, oligomers mobility structure conformation

FCS FCS

C-terminal fusion to GFP or YFP α2AAR-SNAP

agonists-bodipy630 A2AAR-YFP

fusion to GFP, YFP, mCherry antagonist-bodipy630; A1AR-Topaz fusion agonist-bodipy630 A1AR-YFP

FCS FCS

FCS FCS

α1bAR

B2R

labeling antibody-QD antiGFP-biotinstreptavidin-QD655 C-terminal fusion to GFP or YFP

binding, mobility mobility mobility, oligomers mobility mobility, oligomers mobility

FCS

fs-XR FCS

FCS

SPT SPT

mobility mobility, oligomers mobility

A3R

A2AR

A1R

5-HT2BR 5-HT2CR

mobility, oligomers structure mobility

5-HT2AR

question

trafficking mobility

5-HT1AR 5-HT1BR

receptorb

Table 1. Published Single-Molecule Studies on GPCRsa summary

ref

157 626

visualization of ∼140 nm clusters of β1AR and β2AR in live mouse cells unstimulated β1AR and β2AR are confined due to interactions of the C-terminus with PDZ domain and A-kinase anchoring proteins but not caveolae application of PALM to estimate the molecular density in HeLa cells

583

625

623 624

622

620 621

294

618 602 619

607

617

615 616

experimental protocol to study receptor mobility structure with bound antagonist demonstration that pulling on bR in nanodiscs and in native purple membranes yields the same intermediates, demonstrating the usefulness of nanodiscs polypeptide loops potentially act as a barrier to unfolding and contribute significantly to the structural stability of BR individual structural segments of rhodopsin and bR have different properties; a core of rigid structural segments was observed in rhodopsin but not in bR point mutations can reshape the free energy landscape of a membrane protein and force single proteins to populate certain unfolding pathways over others mutations in extracellular Glu and Gln affect unfolding energy landscape characterization of inter- and intramolecular interactions stabilizing structural segments of bR assembled into trimers and dimers, and monomers association of B2R with Gq assessed by FCS; a portion of the receptors diffuses with a diffusion coefficient consistent with dimers or oligomers; no FCCS agonist stimulation increases the lateral mobility of GABAB receptors, but not of β1-/β2ARs

diffusion of Gaq in supported bilayers, either as monomers or as heterotrimers; heterotrimers are more immobile and partition into microdomains near GPCRs brightness analysis shows that the 6 tested receptors diffuse as homodimers in the cell membrane

two populations of diffusing A3AR exist determination of diffusion rate of A3AR dimers, and of the off-rate of the antagonist with and without an allosteric modulator

614

613

agonist-activated receptor is confined when C-terminal cysteine is palmitoylated, explaining the restricted collision coupling to Gs two diffusion states of A2AR in neurons

612 611

610 611

81

608 609

607

571 606

proof-of-concept that these ligands can be used for FCS determination of the diffusion rate of A1AR homodimers, A2AR homodimers, and A1AR−A2AAR heterodimers

uantification of receptor−ligand binding by monitoring the amplitude of the diffusing fraction corresponding to this population in live CHO cells demonstration that fluorescent ligands can be used to monitor diffusion of A1AR in live CHO cells determination of the diffusion rate of A1AR homodimers, A2AR homodimers, and A1AR−A2AAR heterodimers

XFEL structure corresponds to a more accurate room-temperature structure evidence for dimers, without monomers or tetramers

brightness analysis shows that the 6 tested receptors diffuse as homodimers in the cell membrane

internalization and endosomal trafficking of single groups of receptors recruitment of the receptor from soma to dendrites follows an unusual route via vesicle aggregates; lateral diffusion slowed at synapses

Chemical Reviews Review

DOI: 10.1021/acs.chemrev.6b00084 Chem. Rev. XXXX, XXX, XXX−XXX

AI

question

EM SM TIRF, EM AFM

oligomers

structure stoichiometry stability

EP2

mobility, oligomers mobility

SPT

FCS

SMLM

signaling

D1R

colocalization SMLM SM TIRF SMLM

oligomers distribution binding signaling

mCB2R CCR5

SPT

SPT localization SPT

mobility endocytosis mobility

mobility

SM TIRF colocalization FCS, TIRF SPT

stability mobility, oligomers conformation oligomers mobility mobility

AFM FCS

NSOM

distribution distribution structure mobility mobility

NSOM

cAR1-Halo-QD cAR1-eYFP cAR1-eYFP, cAR1-Halo. TMR secondary antibodyQD655 SNAP-505, Halo-TMR mAb-ATTO655 CCL3-Alexa Fluor 647 GFP-nanobodyATTO655 GFP-nanobodyATTO655 C-terminal fusion to GFP or YFP HAtag-AntiHA-QD655

C-terminal fusion to GFP or YFP β2AR-Cy3 SNAP-505, Halo-TMR C5a-YFP cAR1-Halo-TMR

arterenol-Alexa Fluor 532 secondary antibody-Cy2 or Cy3 negative stain β2AR-TMR β2AR-mEos2 β2AR-mEos2 negative stain SNAP-tag secondary antibodyAlexa Fluor 555 β2AR-Venus, β2ARGFP, β2AR-eYFP negative stain β2AR-Cy3, b2AR-Cy5

FCS

labeling TMR TMR SNAP-Alexa Fluor 647

technique

SM confocal SM confocal SPT

EM FCS SMLM SMLM EM SPT SPT

structure

conformation conformation mobility, oligomers mobility, binding oligomers

CB1R

C5a cAR1

receptorb

Table 1. continued

description of the tracking method and labeling protocols

brightness analysis shows that the 6 tested receptors diffuse as homodimers in the cell membrane

303

607

588

305 586 39 70

GABAB2 homodimerizes, β2AR and mCR2 do not dSTORM microscopy of CCR5 in filopodia of CHO cells proof-of-concept ligand-binding assay on CCR5 dSTORM microscopy reveals arrestin3 clustering after stimulation of CCR5 dSTORM microscopy reveals arrestin2 clustering after stimulation of CCR5

633

306 632 303

388 305 631 302

630 607

598 144 629

agonist binding leads to decreased mobility and internalization; decreased mobility key to desensitization

Cy3 used as a probe, fluctuations between two intensity states, active and inactive GABAB2 homodimerizes, β2AR and mCR2 do not kinetics of receptor trapping into clathrin-coated pits cAR1 diffusion is related to microtubule stability but not actin filaments in the amoeba Dictyostelium discoideum; two diffusing populations observed proof of principle of labeling strategy; Halo-QD and Halo-TMR have the same diffusion properties phosphorylation-dependent internalization of cAR1 description of the tracking method and labeling protocols

the Ga-helical domain undergoes a nucleotide- dependent transition from a flexible to a conformationally stabilized state demonstration that β2AR is monomeric in nanodiscs ligand binding induces weak interactions instead of strong localized ones; however, interactions established upon ligand binding are sufficient to change conformational, energetic, kinetic, and mechanical properties of structural segments of β2AR cholesterol increases the kinetic, energetic, and mechanical stability of β2AR brightness analysis shows that the 6 tested receptors diffuse as homodimers in the cell membrane

582

627 387 584 585 599 628 626

157

visualization of ∼140 nm clusters of β1AR and β2AR in live mouse cells description of the methodology observation of conformational substates β2AR partially clusters in cardiomyocytes, but not in other cell lines review (does not contain much information) structure of the receptor−arrestin complex protocol unstimulated β1AR and β2AR are confined due to interactions of the C-terminus with PDZ domain and A-kinase anchoring proteins but not caveolae larger size oligomers observed with GFP and eYFP, whereas size stays constant with Venus upon addition of agonist

594

membrane dynamics and internalization of β2AR in live hippocampal neurons; measurement of KD and kon

ref 27 590 294

summary first observation of the conformational dynamics of a single GPCR follow-up with more experimental details and inverse agonist agonist stimulation increases the lateral mobility of GABAB receptors, but not of β1-/β2ARs

Chemical Reviews Review

DOI: 10.1021/acs.chemrev.6b00084 Chem. Rev. XXXX, XXX, XXX−XXX

SMLM

SPT

mobility

AJ

P2R PAR1

OR17-40

OR5

NPY1R, NPY2R μOR

NK1R

M2R

M1R

LHR

H1R

mobility, oligomers mobility mobility, trafficking mobility binding

FCS

mobility, oligomers mobility, oligomers mobility, signaling mobility mobility, oligomers mobility mobility stoichiometry mobility

labeling

receptors insert into tethered bilayers and aggregate high constitutive activity observed, membrane diffusion and trafficking into endosomes followed

VSVtag-antiVSV-Cy5 OR17-40-GFP

SPT AFM

SPT ATP-QD

receptors insert into tethered bilayers and aggregate

FCS, TIRF

demonstration of labeling method, endocytosis, trafficking, recycling demonstration of method to determine the ligand-binding free-energy landscape

observation of confined diffusion observation of rapid confined diffusion and slow long-range diffusion reconstituted μOR is monomeric; one ligand binds per receptor contrary to Daumas et al.,641 they find rapid hop diffusion

antibody-gold antibody-gold μOR-YFP; agonist-Cy3 antibody-golf, μORmGFP VSVtag-antiVSV-Cy5

SPT SPT TIRF SPT

proof-of-concept labeling, heterogeneous diffusion observed NPY changes mobility and these changes reflect on event related to arrestin recruitment and endocytosis

receptor mobility decreases during the first second after binding of the ligand due to increased interactions with cellular structures

brightness analysis shows that the 6 tested receptors diffuse as homodimers in the cell membrane

brightness analysis shows that the 6 tested receptors diffuse as homodimers in the cell membrane

mobility, clustering, and dimerization kinetics of M1R in CHO cells, randomly distributed with 30% in dimers at any time

super-resolution imaging of functionally asymmetric oligomers reveals diverse functional and structural organizations and the ability to alter signal responses binding of gonadotropin confines the receptor to small domains, maybe rafts

receptor activation increases diffusion; scaffolding protein Homer favors confinement into Homer-mGluR5 clusters after a certain lag time, mGluR5 undergoes directed rearward transport in an actin-dependent way at the surface of neuronal growth cones diffusion of the receptor in the cell membrane

the ligand binding domain exists in three conformations, and orthosteric ligands drive changes between these conformations, leading to dimer interface remodeling mGluR3 has a more stable active state than GluR2 and can be activated by Ca2+

open and closed states of GCGR revealed kinetics of reorientation of extracellular loops; receptors oscillate between a resting and an active conformation on submillisecond time scale

50 647

645 646

644

641 642 573 643

639 640

638

607

607

578

637

587

635 636 593

592

592

600 295

305 628 634

294

agonist stimulation increases the mobility of GABAB receptors, but not of β1-/β2ARs GABAB2 homodimerizes, b2AR and mCR2 do not protocol two populations of GALR observed

304

description of method of observation of dimer lifetimes

ref 579

summary characterization of monomer−dimer equilibrium, unchanged by ligand

NK1R-ACP-CoA-Cy5 receptor-GFP (BiFC)

H1R-YFP, antagonistbodipy630 antiFLAG-Cage500 or Cage552 anti-FLAG-gold (40 nm gold) antagonist-Cy3B, antagonist-AF488 C-terminal fusion to GFP or YFP C-terminal fusion to GFP or YFP NK1R-EGFP

uranyl formate SNAP-mGluR2mGluR2-SNAP SNAP-mGluR2mGluR2-CLIP SNAP-mGluR3mGluR2-CLIP mGluR5-myc-GFP

ligand-Alexa Fluor 594; FPR-YFP (BiFC) Fab-fluorophore, Halotag, ACP-tag SNAP-Alexa Fluor 647 on GABAB1 SNAP-505, Halo-TMR SNAP-tag galanin-rhodamine

SPT FCS

SPT

FCS

SPT

dimers

SPT SPT FCS

mobility mobility binding, mobility oligomers

mGluR5

smFRET

conformation

smFRET

EM smFRET

conformation conformation

conformation

colocalization SPT FCS

mobility, oligomers oligomers mobility mobility

SPT

SPT

oligomers

technique

SPT

question

oligomers

mGluR3

GABAB1GABAB2 GABAB2 GABAB GALR1, GALR2 GCGR mGluR2

FPR

receptorb

Table 1. continued

Chemical Reviews Review

DOI: 10.1021/acs.chemrev.6b00084 Chem. Rev. XXXX, XXX, XXX−XXX

AK

b

a

SPT

TIRF AFM, EM, review AFM AFM AFM AFM AFM

AFM

mobility

mobility

imaging oligomers

stability

FCS, FCCS

SPT

oligomers

mobility

SPT SPT

mobility mobility

EM

structure

AFM

AFM

stability

binding

AFM AFM

stability stability

oligomers oligomers stability stability stability

SPT

structure structure mobility

technique

AFM SPT review EM fs-XR SPT

question

binding mobility

SNAP-tag biotin-tag-streptavidinAlexa Fluor 647 SST-FITC, SSTTexasRed biotin-tag-streptavidinAlexa Fluor 647

negative stain

GTa-peptideATTO647N GTa-peptideATTO647N GTa-peptideATTO647N azF+SpAAC negative stain

gold clusters

ACP-QD

labeling

summary

smoothened and SSTR3 move predominantly by diffusion in cilia; attachment to intraflagellar transport trains is transient and stochastic

homo- and hetero-oligomers are occupied by two ligands

binding events disrupt the primarily diffusive motion of smoothened in cilia smoothened and SSTR3 move predominantly by diffusion in cilia; attachment to intraflagellar transport trains is transient and stochastic

demonstration that rhodopsin forms dimers in native discs rhodopsin and opsin oligomerize in native disc membranes importance of one disulfide bridge for overall stability stabilizing interactions stabilizing mouse and bovine rhodopsin are conserved compared to dark state wild-type rhodopsin, the G90D mutation decreased energetic stability and increased mechanical rigidity of most structural regions in the dark state mutant receptor individual structural segments of rhodopsin and bR have different properties; a core of rigid structural segments was observed in rhodopsin but not in bR Zn2+ increases the stability of most structural segments the absence of palmitoylation in rhodopsin, therefore, destabilizes the molecular interactions formed at the carboxyl terminal end of the receptor, which appears to hinder the activation of transducin by light-activated rhodopsin compared to dark-state rhodopsin, the structural segments stabilizing opsin showed higher interaction strengths and mechanical rigidities and lower conformational variability, lifetimes, and free energies pentameric assembly of the rhodopsin-Gt complex in which a photoactivated rhodopsin dimer serves as a platform for binding the Gt heterotrimer transducer binding establishes localized interactions to tune sensory rhodopsin II

demonstration of labeling, antibody-mediated capturing, and observation on glass surface review on AFM and EM measurements demonstrating oligomeric structure in native membranes

interactions of Gt with rhodopsin are favored at the rim of the membrane

binding of G to rhodopsin, mobility of activated rhodopsin

AFM mapping of two different ligand-binding events using a chemically bifunctionalized AFM tip demonstration of the labeling method fluorescence spectroscopy of rhodopsins demonstration of incorporation of functional rhodopsin into NABBs; EM images demonstrating stoichiometry and orientation active form of rhodopsin bound to arrestin slow and less restricted G diffusion in the active state of rhodopsin

ref

663

664

662 663

661

660

659

657 658

621

155 156 654 655 656

509 580

653

652

648 649 650 145 114 651

This table covers the single-molecule studies on GPCRs to the best of our knowledge. However, we do not guarantee that the list is completely comprehensive and apologize for any unwanted omission. This table only shows the abbreviations of GPCRs. Please refer to Table 2 for their full names.

SSTR1, SSTR5 SSTR3

sensory rhodopsin II smoothened

PTHR rhodopsin

receptorb

Table 1. continued

Chemical Reviews Review

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Chemical Reviews

Review

membrane diffuse into the bleached area. In addition to FRAP, FCS has also been used to investigate the mobility of over 15 different GPCRs in the membrane of live cells.575 However, both FRAP and FCS suffer from the fact that they only yield average diffusion times and can therefore hardly account for local heterogeneities in the cell membrane or provide information on receptor stoichiometry. The single-molecule methods commonly used to study membrane receptor oligomerization include single-molecule photobleaching, smFRET, SMLM, and SPT.576 Single fluorescent molecule video imaging is certainly the most suitable method for visualizing dynamic molecular interactions in live cells and characterizing the diffusion of GPCRs: it not only reports on the variations of the diffusion of fluorescently labeled receptors over time and space, but also tells whether diffusing receptors oligomerize and, if they do, for how long the interaction lasts before an oligomer dissociates into monomers.151,577 The first single-molecule demonstration for the transient dimerization of GPCR in living cells was achieved by Hern et al. utilizing an antagonist of the M1 muscarinic receptor derivatized with Alexa Fluor 488 or Cy3B.578 By tracking individual antagonist-bound receptors in the two channels corresponding to the two fluorophores, the authors were able to demonstrate transient correlated motion of two receptor molecules heterologously expressed in CHO cells, indicating transient dimer formation. They found that the dimers quickly dissociated again into monomers, with an average time constant of 0.7 s (at 23 °C). Whether the M1 muscarinic receptor formed dimers in the absence of the antagonist at physiological temperatures in a more relevant cell line remains unknown. Kasai et al. later fully characterized the monomer−dimer equilibrium of the N-formyl-peptide receptor (FPR) in CHO cells at 37 °C using a fluorescently labeled agonist and an FPRYFP fusion.579 They observed no change in the monomer− dimer equilibrium upon ligand binding, with a typical association time of ∼150 ms and a dissociation time of ∼90 ms. Calebiro et al. observed transient homodimers with lifetimes of about 4 s at 20 °C for β1AR and β2AR labeled via a SNAPtag in CHO cells.294 Similarly, ligand binding did not alter the equilibrium or affect the mobility of the receptors. Interestingly, β2AR seemed to have a higher tendency to form dimers than β1AR at a given expression level. The authors suggest that this apparent difference in converting a collision into a stable interaction might arise from other proteins capable of interfering with dimerization, or from localizations in different microdomains that lead to different effective densities of the two receptors. Calebiro et al. also characterized GABAB receptors for which there is strong evidence for the dimerization between a GABAB1 and a GABAB2 subunit under physiologically relevant conditions. They found GABAB receptors to be in equilibrium between heterodimers and higher-order oligomers, with a preference for tetramers and octamers. With this prototypic class C receptor, an increase in the lateral mobility was observed after agonist binding, suggesting that the ligand can modulate interactions between the receptor and the actin cytoskeleton. All of these studies highlight the dynamic nature of receptor−receptor interactions and suggest that transient dimer or oligomer formation might be a general mechanism for GPCRs, at least in these artificial cellular backgrounds. Kasai et al. even proposed that dynamic homodimers must be crucial for some GPCR functions, which remains to be verified, and

Table 2. Abbreviations of GPCRs GPCR bR 5-HT(x)R A(x)R αxAR βxAR AT1R C5αR cAR1 CB1R CCKAR CCR5 mCBR1 D1R EP2 FPR GABA(x) GCGR mGluR(x) H1R LHR MxR NK(x)R NPY(x)R μOR OR(x) P2R PAR1 PTHR SSTR(x)

G protein-coupled receptor bacteriorhodopsin (not a GPCR, but its heptahelical transmembrane domain is structurally related to GPCRs) 5-hydroxytryptamine (serotonin) receptor subtype x adenosine receptor subtype x αx-adrenergic receptor βx-adrenergic receptor angiotensin II receptor type 1 complement component 5a receptor 1 cAMP receptor 1 cannabinoid type 1 cholecystokinin receptor type A chemokine CCR5 receptor mouse cannabinoid receptor 2 dopamine receptor type 1 prostaglandin E2 receptor N-formyl peptide receptor γ-aminobutyric acid receptor type x glucagon receptor metabotropic glutamate receptor type x histamine receptor type 1 luteinizing hormone receptor muscarinic acetyl choline receptor subtype M-x neurokinin-(x) receptor neuropeptide Y receptor type x μ-opioid receptor olfactory receptor family x P2 purinergic receptor protease-activated receptor-1 parathyroid hormone receptor somatostatin receptor type x

Table 3. Abbreviations of Unnatural Amino Acids uaa 5-OH-Trp AcrK Aladan AlkK AzK Anap AcF azF BCNK BzF (BODIPYFL)K CpK CoK1 CoK2 DansA Ffact HceG NBD-Dap NorK1 NorK2 TCOK ThiPK

unnatural amino acid 5-hydroxyl-L-tryptophan Nε-acryloyl-L-lysine β-[6′-(N,N-dimethyl)amino-2′-naphthoyl]-L-alanine Nε-[(2-propynyloxy)carbonyl]-L-lysine Nε-[(2-azidoethoxy)carbonyl]-L-lysine 3-(6-acetylnaphthalen-2-ylamino)-2-aminopropanoic acid p-acetyl-L-phenylalanine p-azido-L-phenylalanine Nε-[bicyclo[6.1.0]non-4-yn-9-methyloxy)carbonyl]-L-lysine p-benzoyl-L-phenylalanine boron-dipyrromethene-L-lysine Nε-(1-methylcycloprop-2-enecarboxamido)-L-lysine Nε-[(cyclooct-2-yn-1-yloxy)carbonyl]-L-lysine Nε-(2-(cyclooct-2-yn-1-yloxy)ethyl) carbonyl-L-lysine 2-[[5-(dimethylamino)naphthalene-1-yl]sulfonylamino] propanoic acid (dansylalanine) p-fluoroacetyl-L-phenylalanine L-(7-hydroxycoumarin-4-yl)ethylglycine 3-N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl)-2,3diaminopropionic acid Nε-[(5-norbornene-2-yloxy)carbonyl]-L-lysine Nε-[(endonorborn-2-en-5-methyloxy)carbonyl]-L-lysine Nε-[(trans-cyclooctene-4-ol)carbonyl]-L-lysine Nε-thiaprolyl-L-lysine

cell membrane containing the labeled POI is photobleached. The fluorescence intensity then recovers over time because labeled molecules from neighboring regions of the cell AL

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Review

Table 4. Other Abbreviations aaRS ABEL AFM ALEX APD APEX BG BRET BSA BTFA CFP cpGFP CuAAC cryo-EM DEER eDHFR EMCCD EPR FCS FGE FlAsH FP FRAP FRET FTIR GFP GPCR HDX-MS IC ID-PRIME ISC LDAI LDT MAPK mBB (or mBBr) ML

MSD NAMs NSOM PAMs PEG PET PI3K PMMA PMT POE POI PROXYL PSF Pyl ReAsH RhoBo RLuc SAM SCAM sCMOS Sel SILAC smFRET SMLM SpAAC SPIEDAC

aminoacyl tRNA synthetase anti-Brownian electrokinetic atomic force microscopy alternating laser excitation avalanche photodiode ascorbate peroxidase benzylguanine bioluminescence energy transfer bovine serum albumin 3-bromo-1,1,1-trifluoroacetone cyan fluorescent protein circular permuted GFP copper-catalyzed azide−alkyne [3+2] cycloaddition cryo-electron microscopy double electron−electron resonance E. coli dihydrofolate reductase electron-multiplying charge coupled device electron paramagnetic resonance fluorescence correlation spectroscopy formylglycine generating enzyme fluorescein arsenical hairpin binder fluorescent protein fluorescence recovery after photobleaching Förster resonance energy transfer Fourier transformation infrared green fluorescent protein G protein-coupled receptor hydrogen−deuterium exchange mass spectrometry internal conversion interaction-dependent probe incorporation mediated by enzymes intersystem crossing ligand-directed acyl imidazole chemistry ligand-directed tosyl chemistry mitogen-activated protein kinases monobromobimane

SPT TIRF TM TMP TMR Trp Tyr UVB VR YFP

mean squared displacement negative allosteric modulators near-field scanning optical microscopy positive allosteric modulators polyethylene glycol photoinduced electron transfer phosphatidylinositol-3 kinase poly(methyl methacrylate) photomultiplier tube polyoxyethylene protein of interest pyrrolidinyloxy (free radical) point spread function pyrrolysine resorufin arsenical hairpin binder rhodamine-based bisboronic acid Renilla luciferase silent allosteric modulators substituted-cysteine accessibility method scientific complementary metal-oxide-semiconductor selenocysteine stable isotope labeling by amino acids in cell culture single-molecule Förster energy transfer single-molecule localization microscopy strain-promoted azide−alkyne [3+2] cycloaddition strain-promoted inverse-electron-demand Diels−Alder cycloaddition single-particle tracking total-internal-reflection fluorescence transmembrane trimethoprim tetramethylrhodamine tryptophan tyrosine ultraviolet B vibrational relaxation yellow fluorescent protein

maleimide

performed by NSOM using receptors fused to a fluorescent protein. It was found that these receptors were organized in nanodomains with a diameter of ∼150 nm and did not reorganize upon agonist binding.157,582 NSOM is nonetheless particularly difficult to implement in living cells. More recently, SMLM was used to re-evaluate the molecular density of the β2AR in cardiomyocytes, HeLa cells, and CHO cells.583−585 It was found that the receptor indeed preassembled in clusters typically 100−200 nm in size in cardiomyocytes and that the distribution did not significantly change upon addition of ligands, corroborating the findings of the NSOM studies. However, no clustering was observed in HeLa or CHO cells, which was consistent with the findings of the tracking studies discussed earlier294 and with the fact that the β2AR is fully functional as a monomeric entity.144 The absence of significant clustering was further confirmed in CHO and HEK cells by colocalization analysis.305 Similarly, no significant cluster formation in CHO cells was found for the HIV entry coreceptor CCR5, another rhodopsin-like GPCR, although it did accumulate to high densities in the filopodia of these cells.586 On the other hand, the luteinizing hormone receptor (LHR) was shown to mostly form oligomers in HEK cells.587 In this case, however, thanks to an experimental localization

that downstream signaling through G proteins, kinases, or arrestins might be differentially induced by monomers and dimers.577 Single-molecule imaging would be suitable for obtaining a deeper understanding on these biologically relevant questions. 5.2. Membrane Organization beyond the Diffraction Limit

Determining the stoichiometry of higher order oligomers by traditional single-molecule imaging can become very difficult once the number of entities reaches a handful or more. Larger oligomeric assemblies of GPCRs were first described in the context of much debated atomic force microscopy images of rows of rhodopsin dimers in the retina.155,156,580 AFM is however not suitable for most GPCRs, whose expression level under physiological conditions is orders of magnitude lower than rhodopsin. Super-resolution imaging techniques have demonstrated their ability to provide relative or absolute quantitative information about protein copy numbers in oligomers and clusters.69,581 These methods have recently been applied to investigate the organization of GPCRs and arrestins in the cell membrane. Visualizing β1AR and β2AR clusters in cardiomyocytes, whose contraction is controlled by their activation, was first AM

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recording of the fluorescence intensity and lifetime, the authors found a range of discrete conformational states with dwell times of hundreds of milliseconds. The addition of a high affinity agonist increased the dwell times of these states. Millisecond fluctuations were also observed within these conformational states, suggesting that β2AR dynamics spanned a wide range of time scales. Counterintuitively, no large change in the conformational states or the interconversion dynamics occurred upon addition of an agonist, which would be expected from receptor activation. A plausible explanation was the absence of G protein in these experiments. Subsequent structural and NMR studies revealed that a G protein was indispensable for β2AR to reach the fully activated conformational state.221,394 In a recent study, the same β2AR construct was labeled at C265 with Cy3, incorporated into nanodiscs, and immobilized on a glass surface using streptavidin−biotin technology.388 Transitions between two distinct states with dwell times on the order of several seconds were observed in the single-molecule TIRF experiment. The states were assigned to inactive and active-like receptor conformations. Unliganded receptor molecules repeatedly switched between both conformations, leaning toward the inactive conformation. The addition of an agonist favored the active-like conformation, whereas binding of an inverse agonist shifted the conformational distribution toward the inactive conformation. The agonist also enhanced the frequency of activation events, while reducing the frequency of deactivation transitions. The inverse agonist, however, increased the frequency of deactivation events. Aided by molecular modeling, the authors suggested that their ability to observe conformational transitions with Cy3 might have originated in changes in the local confinement of the dye: the inactive conformation was predicted to confine the dye between TM3, 4, and 5, whereas the dye was expected to be in a fully exposed solvent environment in the active state. Cy3 in its lowest excited state returns to the ground state without emission through trans-to-cis isomerization. In the active state of β2AR, the isomerization of Cy3 would be less impeded by the local environment, thereby leading to more efficient quenching of Cy3 fluorescence. This property of Cy3 has been exploited in a variety of studies to probe protein conformational change.591 Experiments relying on a single fluorescent reporter often suffer from an important limitation: signal fluctuation arising from changes in the local environment can barely be correlated to specific structural changes. By comparison, FRET between two fluorophores is more serviceable for interrogating GPCR conformational changes. Previously, FRET experiments on GPCRs have been impeded by the difficulty of attaching synthetic dyes to the receptors. The invention of SNAP- and CLIP-tags has greatly expanded the choices for fluorescent reporters. In a study examining the conformational change of SNAP- and CLIP-tagged metabotropic glutamate receptor (mGluR), which are known to be active only as dimers,295 the kinetics of the reorientation of the extracellular ligand-binding domain of freely diffusing purified receptor dimers were monitored by multiparameter fluorescence detection. In this case, while the observation time of single dimers was limited, the authors reported oscillations between a resting and an active conformation on a submillisecond time scale and showed that agonist efficacy could be correlated with the ability of the ligand to shift the conformational equilibrium toward the active state.

precision of less than 10 nm, analysis of the spatial arrangements of the molecular localization coupled to molecular modeling led the authors to postulate possible structural arrangements for trimers and tetramers. Another question that seems suitable for super-resolution methods to address is the stoichiometry and the duration of the interactions between GPCR and the intracellular adapter proteins. It was recently shown by SMLM imaging of arrestin2 and arrestin3 that different ligands binding to CCR5 induced differential formation of arrestin2 and arrestin3 clusters inside CHO cells.70,588 Whereas most ligands led to recruitment of both arrestins, one ligand caused clustering of arrestin2 but not of arrestin3. In these studies, arrestins were fused to GFP and detected with a fluorescent anti-GFP camel antibody (nanobody). Little is known about the specific physiological roles of arrestin2 and arrestin3, and further investigation with multicolor SMLM is likely to shed some light on the stoichiometry, the duration, and the localization of these interactions. 5.3. Conformational Dynamics

A major motivation for performing experiments on the singlemolecule level is to follow molecular dynamics of unsynchronized molecules in real time with the aim of observing rare or transient states hidden in ensemble measurements.589 Given the relevance of the conformational diversity of GPCRs with respect to their physiological function, it seems crucial to develop a better understanding of the dynamics of GPCR conformations. The development of a conformational assay on the singlemolecule level for GPCRs has nonetheless been hampered by the necessity of finding conditions in which a GPCR could be prepared in a form that is pure enough for single-molecule experiments, functional despite the absence of the cell membrane, and site-specifically labeled in a way that singlemolecule compatible dyes can report the conformational changes. For a long time, the only receptor to meet these very stringent criteria was β2AR, which was specifically labeled on an exposed cysteine residue (C265) in the third intracellular loop by thiol-reactive dyes in the minimal cysteine background.234 Various structurally related ligands were shown to induce different changes in fluorescence intensity of dyes including fluorescein, tetramethylrhodamine (TMR), or bimane specifically attached to C265.102,103,386 Because most of the tested ligands were agonists, such effects could not be unambiguously identified in a functional assay for β2AR that would behave similarly in response to the ligands. The changes in fluorescence intensity were ascribed to changes in the local environment of the dye, most likely related to polarity and confinement. Investigating the dynamics of these conformations was first attempted in an FCS-type of experiment, with single β2AR molecules diffusing through a confocal probe volume.387 The observed histogram of photon counts for single diffusing receptors was broad and displayed two maxima separated by a single bin, possibly corresponding to several conformational states. The major limitation of this study was that the receptors could be observed for no more than a few milliseconds, leaving no time to follow transitions in conformational states. The first observation of a single GPCR changing between its conformations came a decade later.27,590 An ABEL trap allowed β2AR molecules labeled with TMR to be observed for several seconds in detergent micelles. On the basis of simultaneous AN

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binding partners. Over the past decade, structural studies at the single-molecule level have been made possible by atomic force microscopy (AFM), cryo-electron microscopy (cryo-EM), femtosecond serial X-ray crystallography, as well as molecular modeling. Whereas AFM has been primarily used to investigate the mechanical stability of GPCRs in single-molecule force measurements,596 single-particle cryo-EM has revolutionized the field of structural biology in visualizing macromolecular structures, particularly large complexes that have resisted crystallization efforts, at resolutions that can compete against classic crystallography.597 Westfield et al. scrutinized the complex between an agonist, β2AR, and its G protein by cryo-EM.598 They found an overall architecture of the complex in good agreement with the crystal structure of the active-state ternary complex.119 Additionally, they reported the nucleotidedependent rearrangement of α-helical domain of the Gα subunit in the transition from a flexible state to a conformationally stabilized state. In another landmark paper, Shukla et al. examined the interaction between arrestin2 and β2AR by combining singleparticle cryo-EM with hydrogen−deuterium exchange mass spectrometry (HDX-MS), chemical cross-linking, and molecular modeling.599 The authors were able to present the first molecular model of the β2AR-arrestin2 signaling complex by docking the crystal structures of activated arrestin2 and of β2AR into the electron microscopy map densities with constraints provided by HDX-MS and cross-linking, yielding unprecedented insights into the overall architecture of a receptorarrestin complex. More recently, Yang et al. reported the structure of the glucagon receptor (GCGR) by using the same combination of techniques.600 This study revealed the open and closed states of GCGR, suggesting the glucagon binding through a conformational selection mechanism. Obtaining full-length structures at an atomic resolution of GCGR, a class B GPCR, which does not belong to the most abundant class-A (rhodopsin-like) GPCR family, has been very challenging due to the very dynamic nature of their extracellular domain and lack of highaffinity ligands to stabilize the receptor structure. The approach based on single-particle cryo-EM therefore appears very promising in complementing traditional crystallographic methods to provide insights into the structure and dynamics of GPCRs. In parallel to EM, breakthroughs in GPCR structural studies can be expected in the near future from pump−probe serial femtosecond crystallography, which uses the potential of X-ray free electron lasers for tracking the dynamics of light-triggered processes, such as rhodopsin activation.601 The static roomtemperature structures of an antagonist-bound angiotensin receptor602 and of rhodopsin bound to visual arrestin114 were solved by serial femtosecond X-ray crystallography. The possibility of obtaining meaningful static and dynamic structural information by molecular modeling and molecular dynamics simulations deserves to be credited here. With the ever increasing computational power of supercomputers, simulations spanning tens of microseconds of dynamics can be robustly performed to evaluate processes ranging from the activation mechanism of GPCRs to nucleotide exchange in heterotrimeric G proteins,603,604 demonstrating the potential of this type of simulations to serve as a “computational microscope”.605

In a follow-up study, Isacoff and co-workers harnessed the full power of single-molecule FRET to probe the conformational reorientation of the ligand-binding domain of mGluR homodimers.592 SNAP−mGluR and CLIP−mGluR were expressed in HEK293T cells, labeled a FRET donor and acceptor fluorophore, respectively, purified from the cell membrane, and captured on a glass surface in the scheme shown in Figure 6. Single dimers could be continuously observed for several tens of seconds. The authors were able to demonstrate that the ligand-binding domains interconverted between three conformations: resting, activated, and a shortlived intermediate state. Agonists induced transitions between these conformational states, with their efficacies being determined by occupancy of the active conformation. Overall, their results support a general mechanism for the activation of mGluR: agonist binding induces closure of the ligand-binding domain, followed by dimer interface reorientation. 5.4. Ligand Binding

Ligand binding to its GPCR is the key molecular event for triggering an intracellular signaling response. Nonetheless, literature reporting on ligand binding at the single molecule level is scarce. Such studies are highly desirable because they can provide quantitative information about the interaction between two molecules for deriving kinetic (kon and koff) and thermodynamic parameters (the equilibrium constant KD = koff/kon). A possible reason is that the accessible concentration range of single-molecule experiments typically falls into the picomolar to nanomolar range, whereas many biomolecular interactions require concentrations at least 100 times larger.18 Furthermore, a single-molecule binding assay demands both the receptor and the ligand to be simultaneously monitored; thus a fluorescent tag also needs to be attached to the ligand without impairing the receptor−ligand interaction. Whereas coupling a fluorescent tag to a peptide ligand without affecting its binding property seems reasonably easy if the peptide is large enough, this task proves much more complicated for small-molecule ligands.202,204,217 Fluorescent ligands have therefore often been used as an indirect way of labeling the receptor, for example, to follow the lateral mobility of the latter in the plasma membrane of living cells. Importantly, they increasingly replace dangerous radioactive tracers in affinity measurements based on integration of the total fluorescence response over a whole field of view.593 Despite all of these difficulties, FCS was used in one case to determine the binding constant of the fluorescently labeled agonist arterenol to β2AR in hippocampal neurons and in an epithelial cell line, with the KD determined to be 1.3 and 6.0 nM, respectively.594 The feasibility of a single-molecule binding assay on GPCRs was later evaluated for the receptor CCR5, whose natural ligands chemokines are ∼70 amino acid long peptides. The receptor was purified and embedded in a membrane tethered to the surface of a chip. Binding events of the fluorescently labeled chemokine CCL3 with CCR5, whose residence time lasted tens of minutes, could be monitored over the time course of hours.39 5.5. Structure and Stability

A detailed understanding of macromolecular processes and their dynamics requires the integration of information from a wide range of experimental and computational approaches covering different spatial and temporal regimes.595 Fluorescence methods can be limited when it comes to obtaining highresolution structural information for the receptor and its AO

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6. CONCLUSION AND PROSPECT Biochemical and biophysical characterization of GPCRs in the past two decades has led to a considerable appreciation of the dynamic and heterogeneous nature of GPCR signaling complex, which has given rise to the increasing popularity of single-molecule techniques in the GPCR field. The singlemolecule studies of GPCRs have benefitted from the interdisciplinary efforts in three areas. First, an intimate knowledge of the receptor biochemistry and cell biology necessarily underlies the success of any experiment. Thanks to the explosive development of molecular cloning and sequencing, experiments on heterologously expressed GPCRs in living cells have become more tractable. By comparison, studying purified GPCRs in a reconstituted system, despite its longer history, remains a major challenge. As a result, the literature of the biochemical studies on two prototypical GPCRs, rhodopsin and β2AR, appears to be disproportionally rich. Second, innovative single-molecule detection schemes and instrumentations have enabled a broader range of questions to be approached. Finally, the expanding chemical biology toolkits for protein modification have been harnessed to prepare labeled receptors that suit specific experimental designs. Labeling strategies aiming at facilitating single-molecule studies should involve receptors with minimal alterations of their native structure. The possibility of targeting GPCRs in native tissues with fluorescent ligands or ligand-directed labeling may overcome the limitations associated with heterologous expression systems, such as overexpression of receptor or cell type-specific background. Chemoenzymatic labeling provides a general approach for tagging GPCRs with different types of fluorescent labels. Previously, the transmembrane region and the ligand binding pockets of GPCRs were mostly studied by site-directed mutagenesis. The application of unnatural amino acids, combined with bioorthogonal labeling chemistries, may significantly advance spectroscopic and microscopic characterization of receptor conformational change.

photochemistry and ultrafast dynamics of biomolecules with Eric Vauthey at the University of Geneva (Ph.D. 2007) before moving into single-molecule spectroscopy and imaging as a postdoc with W. E. Moerner at Stanford University (2008−2010). In 2010, he started his independent research thanks to an Ambizione fellowship of the Swiss National Science Foundation at the Faculty of Medicine of the University of Geneva, his research focusing on the development and application of single-molecule tools for biology with an emphasis on the dynamics of G protein-coupled receptors. He has been a visiting assistant professor at The Rockefeller University in the group of Thomas P. Sakmar since October 2015. Thomas Huber graduated from the University of Munich in Medicine in 1995. He conduced his Ph.D. work with Klaus Beyer on NMR spectroscopy and molecular dynamics simulations of biological membranes in Martin Klingenberg’s Institute of Physical Biochemistry at the University of Munich (Ph.D. 1999). Huber then performed postdoctoral work with Michael F. Brown in the Department of Chemistry at the University of Arizona in Tucson to study lipid− protein interactions. He joined Thomas P. Sakmar’s laboratory at the Rockefeller University in 2002. Here, he studied receptor oligomerization and developed applications of unnatural amino acid mutagenesis in GPCR drug discovery. In 2013, he was appointed Research Assistant Professor. His research interests are in the area of Chemical and Quantitative Biology with a focus on single-molecule methods.

ACKNOWLEDGMENTS We acknowledge the generous support from a grant from the Robertson Foundation, the Crowley Family Fund, the Danica Foundation, and the NIH R01 EY012049 to T.H., as well as the Tri-Institutional Training Program in Chemical Biology for supporting H.T. This work was also generously supported by an International Research Alliance with Thue W. Schwartz at The Novo Nordisk Foundation Center for Basic Metabolic Research (http://www.metabol.ku.dk) through an unconditional grant from the Novo Nordisk Foundation to the University of Copenhagen. We also acknowledge Thomas P. Sakmar and the following members of his lab who contributed to this work: Kelly Daggett, Amy Grunbeck, Manija Kazmi, Adam Knepp, Saranga Naganathan, Minyoung Park, Sarmistha Ray-Saha, Carlos Rico, Pallavi Sachdev, Louise ValentinHansen, and Shixin Ye.

AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected] Author Contributions †

H.T. and A.F. contributed equally to this work.

Notes

REFERENCES

The authors declare no competing financial interest.

(1) Overington, J. P.; Al-Lazikani, B.; Hopkins, A. L. How Many Drug Targets Are There? Nat. Rev. Drug Discovery 2006, 5, 993−996. (2) Rask-Andersen, M.; Almen, M. S.; Schioth, H. B. Trends in the Exploitation of Novel Drug Targets. Nat. Rev. Drug Discovery 2011, 10, 579−590. (3) Kobilka, B. The Structural Basis of G-Protein-Coupled Receptor Signaling (Nobel Lecture). Angew. Chem., Int. Ed. 2013, 52, 6380− 6388. (4) Köhrer, C.; Sullivan, E. L.; RajBhandary, U. L. Complete Set of Orthogonal 21st Aminoacyl-tRNA Synthetase-Amber, Ochre and Opal Suppressor tRNA Pairs: Concomitant Suppression of Three Different Termination Codons in an mRNA in Mammalian Cells. Nucleic Acids Res. 2004, 32, 6200−6211. (5) Stevens, R. C.; Cherezov, V.; Katritch, V.; Abagyan, R.; Kuhn, P.; Rosen, H.; Wuthrich, K. The GPCR Network: A Large-Scale Collaboration to Determine Human GPCR Structure and Function. Nat. Rev. Drug Discovery 2012, 12, 25−34. (6) Moerner, W. E.; Kador, L. Optical Detection and Spectroscopy of Single Molecules in a Solid. Phys. Rev. Lett. 1989, 62, 2535−2538.

Biographies He Tian received her B.Sc. in Chemistry from Peking University, China, in 2009. She then moved to New York City to enroll in the TriInstitutional Ph.D. Program in Chemical Biology. She conducted her doctoral research under the supervision of Thomas P. Sakmar and Thomas Huber at the Rockefeller University and obtained her Ph.D. in 2015. Her dissertation focused on developing chemical biology tools for probing the structure−function relationship in G protein-coupled receptors. In 2016, she joined the research group led by Adam Cohen in the Department of Chemistry and Chemical Biology at Harvard University as a postdoctoral researcher. Her current research interest involves understanding the biophysical principles governing membrane proteins by protein engineering. Alexandre Fürstenberg studied chemistry and biochemistry at the Universities of Lausanne and Geneva in Switzerland. He specialized in AP

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(7) Shera, E. B.; Seitzinger, N. K.; Davis, L. M.; Keller, R. A.; Soper, S. A. Detection of Single Fluorescent Molecules. Chem. Phys. Lett. 1990, 174, 553−557. (8) Joo, C.; Balci, H.; Ishitsuka, Y.; Buranachai, C.; Ha, T. Advances in Single-Molecule Fluorescence Methods for Molecular Biology. Annu. Rev. Biochem. 2008, 77, 51−76. (9) Moerner, W. E.; Shechtman, Y.; Wang, Q. Single-Molecule Spectroscopy and Imaging over the Decades. Faraday Discuss. 2015, 184, 9−36. (10) Diez, M.; Zimmermann, B.; Borsch, M.; Konig, M.; Schweinberger, E.; Steigmiller, S.; Reuter, R.; Felekyan, S.; Kudryavtsev, V.; Seidel, C. A. M.; et al. Proton-Powered Subunit Rotation in Single Membrane-Bound F0F1-Atp Synthase. Nat. Struct. Mol. Biol. 2004, 11, 135−141. (11) Uemura, S.; Aitken, C. E.; Korlach, J.; Flusberg, B. A.; Turner, S. W.; Puglisi, J. D. Real-Time tRNA Transit on Single Translating Ribosomes at Codon Resolution. Nature 2010, 464, 1012−1017. (12) Ambrose, W. P.; Moerner, W. E. Fluorescence Spectroscopy and Spectral Diffusion of Single Impurity Molecules in a Crystal. Nature 1991, 349, 225−227. (13) Trautman, J. K.; Macklin, J. J.; Brus, L. E.; Betzig, E. Near-Field Spectroscopy of Single Molecules at Room-Temperature. Nature 1994, 369, 40−42. (14) Wang, Q.; Moerner, W. E. Lifetime and Spectrally Resolved Characterization of the Photodynamics of Single Fluorophores in Solution Using the Anti-Brownian Electrokinetic Trap. J. Phys. Chem. B 2013, 117, 4641−4648. (15) Moerner, W. E.; Orrit, M. Illuminating Single Molecules in Condensed Matter. Science 1999, 283, 1670−1676. (16) Weiss, S. Fluorescence Spectroscopy of Single Biomolecules. Science 1999, 283, 1676−1683. (17) Moerner, W. E.; Fromm, D. P. Methods of Single-Molecule Fluorescence Spectroscopy and Microscopy. Rev. Sci. Instrum. 2003, 74, 3597−3619. (18) Holzmeister, P.; Acuna, G. P.; Grohmann, D.; Tinnefeld, P. Breaking the Concentration Limit of Optical Single-Molecule Detection. Chem. Soc. Rev. 2014, 43, 1014−1028. (19) Moerner, W. E. New Directions in Single-Molecule Imaging and Analysis. Proc. Natl. Acad. Sci. U. S. A. 2007, 104, 12596−12602. (20) Ha, T.; Tinnefeld, P. Photophysics of Fluorescent Probes for Single-Molecule Biophysics and Super-Resolution Imaging. Annu. Rev. Phys. Chem. 2012, 63, 595−617. (21) Juette, M. F.; Terry, D. S.; Wasserman, M. R.; Zhou, Z.; Altman, R. B.; Zheng, Q.; Blanchard, S. C. The Bright Future of SingleMolecule Fluorescence Imaging. Curr. Opin. Chem. Biol. 2014, 20, 103−111. (22) Michalet, X.; Weiss, S.; Jager, M. Single-Molecule Fluorescence Studies of Protein Folding and Conformational Dynamics. Chem. Rev. 2006, 106, 1785−1813. (23) Lord, S. J.; Lee, H. L. D.; Moerner, W. E. Single-Molecule Spectroscopy and Imaging of Biomolecules in Living Cells. Anal. Chem. 2010, 82, 2192−2203. (24) Ulbrich, M. H.; Isacoff, E. Y. Subunit Counting in MembraneBound Proteins. Nat. Methods 2007, 4, 319−321. (25) Jiang, Y.; Douglas, N. R.; Conley, N. R.; Miller, E. J.; Frydman, J.; Moerner, W. E. Sensing Cooperativity in ATP Hydrolysis for Single Multisubunit Enzymes in Solution. Proc. Natl. Acad. Sci. U. S. A. 2011, 108, 16962−16967. (26) Kühnemuth, R.; Seidel, C. A. M. Principles of Single Molecule Multiparameter Fluorescence Spectroscopy. Single Mol. 2001, 2, 251− 254. (27) Bockenhauer, S.; Fürstenberg, A.; Yao, X. J.; Kobilka, B. K.; Moerner, W. E. Conformational Dynamics of Single G ProteinCoupled Receptors in Solution. J. Phys. Chem. B 2011, 115, 13328− 13338. (28) Marme, N.; Knemeyer, J. P.; Sauer, M.; Wolfrum, J. Inter- and Intramolecular Fluorescence Quenching of Organic Dyes by Tryptophan. Bioconjugate Chem. 2003, 14, 1133−1139.

(29) Doose, S.; Neuweiler, H.; Sauer, M. A Close Look at Fluorescence Quenching of Organic Dyes by Tryptophan. ChemPhysChem 2005, 6, 2277−2285. (30) Doose, S.; Neuweiler, H.; Sauer, M. Fluorescence Quenching by Photoinduced Electron Transfer: A Reporter for Conformational Dynamics of Macromolecules. ChemPhysChem 2009, 10, 1389−1398. (31) Yang, H.; Luo, G. B.; Karnchanaphanurach, P.; Louie, T. M.; Rech, I.; Cova, S.; Xun, L. Y.; Xie, X. S. Protein Conformational Dynamics Probed by Single-Molecule Electron Transfer. Science 2003, 302, 262−266. (32) Haustein, E.; Schwille, P. Fluorescence Correlation Spectroscopy: Novel Variations of an Established Technique. Annu. Rev. Biophys. Biomol. Struct. 2007, 36, 151−169. (33) Saxton, M. J.; Jacobson, K. Single-Particle Tracking: Applications to Membrane Dynamics. Annu. Rev. Biophys. Biomol. Struct. 1997, 26, 373−399. (34) Dahan, M.; Levi, S.; Luccardini, C.; Rostaing, P.; Riveau, B.; Triller, A. Diffusion Dynamics of Glycine Receptors Revealed by Single-Quantum Dot Tracking. Science 2003, 302, 442−445. (35) Roy, R.; Hohng, S.; Ha, T. A Practical Guide to Single-Molecule FRET. Nat. Methods 2008, 5, 507−516. (36) Bartko, A. P.; Dickson, R. M. Imaging Three-Dimensional Single Molecule Orientations. J. Phys. Chem. B 1999, 103, 11237−11241. (37) Willets, K. A.; Ostroverkhova, O.; He, M.; Twieg, R. J.; Moerner, W. E. Fluorophores for Single-Molecule Imaging. J. Am. Chem. Soc. 2003, 125, 1174−1175. (38) Jeyachandran, Y. L.; Mielczarski, J. A.; Mielczarski, E.; Rai, B. Efficiency of Blocking of Non-Specific Interaction of Different Proteins by BSA Adsorbed on Hydrophobic and Hydrophilic Surfaces. J. Colloid Interface Sci. 2010, 341, 136−142. (39) Huber, T.; Sakmar, T. P. New Approaches for Studying the Dynamic Assembly and Activation of GPCR Signaling Complexes. Trends Pharmacol. Sci. 2011, 32, 410−419. (40) Jain, A.; Liu, R.; Ramani, B.; Arauz, E.; Ishitsuka, Y.; Ragunathan, K.; Park, J.; Chen, J.; Xiang, Y. K.; Ha, T. Probing Cellular Protein Complexes Using Single-Molecule Pull-Down. Nature 2011, 473, 484−488. (41) Yeom, K. H.; Heo, I.; Lee, J.; Hohng, S.; Kim, V. N.; Joo, C. Single-Molecule Approach to Immunoprecipitated Protein Complexes: Insights into miRNA Uridylation. EMBO Rep. 2011, 12, 690−696. (42) Sofia, S. J.; Premnath, V. V.; Merrill, E. W. Poly(ethylene oxide) Grafted to Silicon Surfaces: Grafting Density and Protein Adsorption. Macromolecules 1998, 31, 5059−5070. (43) Chandradoss, S. D.; Haagsma, A. C.; Lee, Y. K.; Hwang, J. H.; Nam, J. M.; Joo, C. Surface Passivation for Single-molecule Protein Studies. J. Vis. Exp. 2014, DOI: 10.3791/50549. (44) Hua, B.; Han, K. Y.; Zhou, R.; Kim, H.; Shi, X.; Abeysirigunawardena, S. C.; Jain, A.; Singh, D.; Aggarwal, V.; Woodson, S. A.; et al. An Improved Surface Passivation Method for Single-Molecule Studies. Nat. Methods 2014, 11, 1233−1236. (45) Cohen, A. E.; Moerner, W. E. Suppressing Brownian Motion of Individual Biomolecules in Solution. Proc. Natl. Acad. Sci. U. S. A. 2006, 103, 4362−4365. (46) Cohen, A. E.; Moerner, W. E. Controlling Brownian Motion of Single Protein Molecules and Single Fluorophores in Aqueous Buffer. Opt. Express 2008, 16, 6941−6956. (47) Fields, A. P.; Cohen, A. E. Electrokinetic Trapping at the One Nanometer Limit. Proc. Natl. Acad. Sci. U. S. A. 2011, 108, 8937−8942. (48) Wang, Q.; Goldsmith, R. H.; Jiang, Y.; Bockenhauer, S. D.; Moerner, W. E. Probing Single Biomolecules in Solution Using the Anti-Brownian Electrokinetic (ABEL) Trap. Acc. Chem. Res. 2012, 45, 1955−1964. (49) Cohen, A. E.; Moerner, W. E. Principal-Components Analysis of Shape Fluctuations of Single DNA Molecules. Proc. Natl. Acad. Sci. U. S. A. 2007, 104, 12622−12627. (50) Jiang, S.; Liu, A.; Duan, H.; Soo, J.; Chen, P. Labeling and Tracking P2 Purinergic Receptors in Living Cells Using ATPConjugated Quantum Dots. Adv. Funct. Mater. 2011, 21, 2776−2780. AQ

DOI: 10.1021/acs.chemrev.6b00084 Chem. Rev. XXXX, XXX, XXX−XXX

Chemical Reviews

Review

(51) Goldsmith, R. H.; Moerner, W. E. Watching Conformationaland Photodynamics of Single Fluorescent Proteins in Solution. Nat. Chem. 2010, 2, 179−186. (52) Betzig, E.; Chichester, R. J. Single Molecules Observed by NearField Scanning Optical Microscopy. Science 1993, 262, 1422−1425. (53) Betzig, E. Single Molecules, Cells, and Super-Resolution Optics (Nobel Lecture). Angew. Chem., Int. Ed. 2015, 54, 8034−8053. (54) Betzig, E. Proposed Method for Molecular Optical Imaging. Opt. Lett. 1995, 20, 237−239. (55) Hell, S. W.; Wichmann, J. Breaking the Diffraction Resolution Limit by Stimulated-Emission - Stimulated-Emission-Depletion Fluorescence Microscopy. Opt. Lett. 1994, 19, 780−782. (56) Hell, S. W. Far-Field Optical Nanoscopy. Science 2007, 316, 1153−1158. (57) Allen, J. R.; Ross, S. T.; Davidson, M. W. Structured Illumination Microscopy for Superresolution. ChemPhysChem 2014, 15, 566−576. (58) Betzig, E.; Patterson, G. H.; Sougrat, R.; Lindwasser, O. W.; Olenych, S.; Bonifacino, J. S.; Davidson, M. W.; Lippincott-Schwartz, J.; Hess, H. F. Imaging Intracellular Fluorescent Proteins at Nanometer Resolution. Science 2006, 313, 1642−1645. (59) Hess, S. T.; Girirajan, T. P. K.; Mason, M. D. Ultra-High Resolution Imaging by Fluorescence Photoactivation Localization Microscopy. Biophys. J. 2006, 91, 4258−4272. (60) Rust, M. J.; Bates, M.; Zhuang, X. W. Sub-Diffraction-Limit Imaging by Stochastic Optical Reconstruction Microscopy (STORM). Nat. Methods 2006, 3, 793−795. (61) Heilemann, M.; van de Linde, S.; Schuttpelz, M.; Kasper, R.; Seefeldt, B.; Mukherjee, A.; Tinnefeld, P.; Sauer, M. SubdiffractionResolution Fluorescence Imaging with Conventional Fluorescent Probes. Angew. Chem., Int. Ed. 2008, 47, 6172−6176. (62) Fürstenberg, A.; Heilemann, M. Single-Molecule Localization Microscopy - Near-Molecular Spatial Resolution in Light Microscopy with Photoswitchable Fluorophores. Phys. Chem. Chem. Phys. 2013, 15, 14919−14930. (63) Yildiz, A.; Selvin, P. R. Fluorescence Imaging with One Manometer Accuracy: Application to Molecular Motors. Acc. Chem. Res. 2005, 38, 574−582. (64) Moerner, W. E. Single-Molecule Mountains Yield Nanoscale Cell Images. Nat. Methods 2006, 3, 781−782. (65) Moerner, W. E. Microscopy beyond the Diffraction Limit Using Actively Controlled Single Molecules. J. Microsc. 2012, 246, 213−220. (66) Klein, T.; Proppert, S.; Sauer, M. Eight Years of Single-Molecule Localization Microscopy. Histochem. Cell Biol. 2014, 141, 561−575. (67) Patterson, G.; Davidson, M.; Manley, S.; Lippincott-Schwartz, J. Superresolution Imaging using Single-Molecule Localization. Annu. Rev. Phys. Chem. 2010, 61, 345−367. (68) Gahlmann, A.; Moerner, W. E. Exploring Bacterial Cell Biology with Single-Molecule Tracking and Super-Resolution Imaging. Nat. Rev. Microbiol. 2013, 12, 9−22. (69) Fricke, F.; Beaudouin, J.; Eils, R.; Heilemann, M. One, Two or Three? Probing the Stoichiometry of Membrane Proteins by SingleMolecule Localization Microscopy. Sci. Rep. 2015, 5, 14072. (70) Tarancon Diez, L.; Bönsch, C.; Malkusch, S.; Truan, Z.; Munteanu, M.; Heilemann, M.; Hartley, O.; Endesfelder, U.; Fürstenberg, A. Coordinate-Based Co-Localization-Mediated Analysis of Arrestin Clustering Upon Stimulation of the C-C Chemokine Receptor 5 with Rantes/CCL5 Analogues. Histochem. Cell Biol. 2014, 142, 69−77. (71) Churchman, L. S.; Okten, Z.; Rock, R. S.; Dawson, J. F.; Spudich, J. A. Single Molecule High-Resolution Colocalization of Cy3 and Cy5 Attached to Macromolecules Measures Intramolecular Distances through Time. Proc. Natl. Acad. Sci. U. S. A. 2005, 102, 1419−1423. (72) Malkusch, S.; Endesfelder, U.; Mondry, J.; Gelleri, M.; Verveer, P. J.; Heilemann, M. Coordinate-Based Colocalization Analysis of Single-Molecule Localization Microscopy Data. Histochem. Cell Biol. 2012, 137, 1−10.

(73) Ha, T.; Enderle, T.; Ogletree, D. F.; Chemla, D. S.; Selvin, P. R.; Weiss, S. Probing the Interaction between Two Single Molecules: Fluorescence Resonance Energy Transfer between a Single Donor and a Single Acceptor. Proc. Natl. Acad. Sci. U. S. A. 1996, 93, 6264−6268. (74) Kapanidis, A. N.; Lee, N. K.; Laurence, T. A.; Doose, S.; Margeat, E.; Weiss, S. Fluorescence-Aided Molecule Sorting: Analysis of Structure and Interactions by Alternating-Laser Excitation of Single Molecules. Proc. Natl. Acad. Sci. U. S. A. 2004, 101, 8936−8941. (75) Kapanidis, A. N.; Laurence, T. A.; Lee, N. K.; Margeat, E.; Kong, X. X.; Weiss, S. Alternating-Laser Excitation of Single Molecules. Acc. Chem. Res. 2005, 38, 523−533. (76) Santoso, Y.; Joyce, C. M.; Potapova, O.; Le Reste, L.; Hohlbein, J.; Torella, J. P.; Grindley, N. D. F.; Kapanidis, A. N. Conformational Transitions in DNA Polymerase I Revealed by Single-Molecule FRET. Proc. Natl. Acad. Sci. U. S. A. 2010, 107, 715−720. (77) Munro, J. B.; Wasserman, M. R.; Altman, R. B.; Wang, L.; Blanchard, S. C. Correlated Conformational Events in EF-G and the Ribosome Regulate Translocation. Nat. Struct. Mol. Biol. 2010, 17, 1470−1477. (78) Lefkowitz, R. J. A Brief History of G Protein-Coupled Receptors (Nobel Lecture). Angew. Chem., Int. Ed. 2013, 52, 6366−6378. (79) Bockaert, J.; Pin, J. P. Molecular Tinkering of G ProteinCoupled Receptors: An Evolutionary Success. EMBO J. 1999, 18, 1723−1729. (80) Lagerstrom, M. C.; Schioth, H. B. Structural Diversity of G Protein-Coupled Receptors and Significance for Drug Discovery. Nat. Rev. Drug Discovery 2008, 7, 339−357. (81) Briddon, S. J.; Middleton, R. J.; Cordeaux, Y.; Flavin, F. M.; Weinstein, J. A.; George, M. W.; Kellam, B.; Hill, S. J. Quantitative Analysis of the Formation and Diffusion of A(1)-Adenosine ReceptorAntagonist Complexes in Single Living Cells. Proc. Natl. Acad. Sci. U. S. A. 2004, 101, 4673−4678. (82) Oldham, W. M.; Hamm, H. E. Heterotrimeric G Protein Activation by G-Protein-Coupled Receptors. Nat. Rev. Mol. Cell Biol. 2008, 9, 60−71. (83) Sun, Y.; McGarrigle, D.; Huang, X. Y. When a G ProteinCoupled Receptor Does Not Couple to a G Protein. Mol. BioSyst. 2007, 3, 849−854. (84) Moore, C. A. C.; Milano, S. K.; Benovic, J. L. Regulation of Receptor Trafficking by GRKs and Arrestins. Annu. Rev. Physiol. 2007, 69, 451−482. (85) Hanyaloglu, A. C.; von Zastrow, M. Regulation of GPCRs by Endocytic Membrane Trafficking and Its Potential Implications. Annu. Rev. Pharmacol. Toxicol. 2008, 48, 537−568. (86) DeWire, S. M.; Ahn, S.; Lefkowitz, R. J.; Shenoy, S. K. βArrestins and Cell Signaling. Annu. Rev. Physiol. 2007, 69, 483−510. (87) Kühne, W. On the Photochemistry in the Retina and on Visual Purple; MacMillan and Co.: London, 1878. (88) Bennett, M. R. One Hundred Years of Adrenaline: the Discovery of Autoreceptors. Clin. Auton. Res. 1999, 9, 145−159. (89) Rubin, R. P. A Brief History of Great Discoveries in Pharmacology: In Celebration of the Centennial Anniversary of the Founding of the American Society of Pharmacology and Experimental Therapeutics. Pharmacol. Rev. 2007, 59, 289−359. (90) Dixon, R. A. F.; Kobilka, B. K.; Strader, D. J.; Benovic, J. L.; Dohlman, H. G.; Frielle, T.; Bolanowski, M. A.; Bennett, C. D.; Rands, E.; Diehl, R. E.; et al. Cloning of the Gene and Cdna for Mammalian β-Adrenergic-Receptor and Homology with Rhodopsin. Nature 1986, 321, 75−79. (91) Kobilka, B. K.; Matsui, H.; Kobilka, T. S.; Yang-Feng, T. L.; Francke, U.; Caron, M. G.; Lefkowitz, R. J.; Regan, J. W. Cloning, Sequencing, and Expression of the Gene Coding for the Human Platelet α2-Adrenergic Receptor. Science 1987, 238, 650−656. (92) Frielle, T.; Collins, S.; Daniel, K. W.; Caron, M. G.; Lefkowitz, R. J.; Kobilka, B. K. Cloning of the cDNA for the Human β1Adrenergic Receptor. Proc. Natl. Acad. Sci. U. S. A. 1987, 84, 7920− 7924. (93) Bitensky, M. W.; Wheeler, M. A.; Rasenick, M. M.; Yamazaki, A.; Stein, P. J.; Halliday, K. R.; Wheeler, G. L. Functional Exchange of AR

DOI: 10.1021/acs.chemrev.6b00084 Chem. Rev. XXXX, XXX, XXX−XXX

Chemical Reviews

Review

Components between Light-Activated Photoreceptor Phosphodiesterase and Hormone-Activated Adenylate-Cyclase Systems. Proc. Natl. Acad. Sci. U. S. A. 1982, 79, 3408−3412. (94) Stryer, L. Transducin and the Cyclic-GMP Phosphodiesterase Amplifier Proteins in Vision. Cold Spring Harbor Symp. Quant. Biol. 1983, 48, 841−852. (95) Stryer, L. Cyclic-GMP Cascade of Vision. Annu. Rev. Neurosci. 1986, 9, 87−119. (96) Dohlman, H. G.; Caron, M. G.; Lefkowitz, R. J. A Family of Receptors Coupled to Guanine-Nucleotide Regulatory Proteins. Biochemistry 1987, 26, 2657−2664. (97) Kobilka, B. K. Amino and Carboxyl Terminal Modifications to Facilitate the Production and Purification of a G Protein-Coupled Receptor. Anal. Biochem. 1995, 231, 269−271. (98) Farrens, D. L.; Altenbach, C.; Yang, K.; Hubbell, W. L.; Khorana, H. G. Requirement of Rigid-Body Motion of Transmembrane Helices for Light Activation of Rhodopsin. Science 1996, 274, 768−770. (99) Sheikh, S. P.; Zvyaga, T. A.; Lichtarge, O.; Sakmar, T. P.; Bourne, H. R. Rhodopsin Activation Blocked by Metal-Ion-Binding Sites Linking Transmembrane Helices C and F. Nature 1996, 383, 347−350. (100) Gether, U.; Lin, S.; Ghanouni, P.; Ballesteros, J. A.; Weinstein, H.; Kobilka, B. K. Agonists Induce Conformational Changes in Transmembrane Domains III and VI of the β(2) Adrenoceptor. EMBO J. 1997, 16, 6737−6747. (101) Jensen, A. D.; Guarnieri, F.; Rasmussen, S. G. F.; Asmar, F.; Ballesteros, J. A.; Gether, U. Agonist-Induced Conformational Changes at the Cytoplasmic Side of Transmembrane Segment 6 in the β(2) Adrenergic Receptor Mapped by Site-Selective Fluorescent Labeling. J. Biol. Chem. 2001, 276, 9279−9290. (102) Ghanouni, P.; Gryczynski, Z.; Steenhuis, J. J.; Lee, T. W.; Farrens, D. L.; Lakowicz, J. R.; Kobilka, B. K. Functionally Different Agonists Induce Distinct Conformations in the G Protein Coupling Domain of the β(2) Adrenergic Receptor. J. Biol. Chem. 2001, 276, 24433−24436. (103) Swaminath, G.; Xiang, Y.; Lee, T. W.; Steenhuis, J.; Parnot, C.; Kobilka, B. K. Sequential Binding of Agonists to the β(2) Adrenoceptor: Kinetic Evidence for Intermediate Conformational States. J. Biol. Chem. 2004, 279, 686−691. (104) Swaminath, G.; Deupi, X.; Lee, T. W.; Zhu, W.; Thian, F. S.; Kobilka, T. S.; Kobilka, B. Probing the β(2) Adrenoceptor Binding Site with Catechol Reveals Differences in Binding and Activation by Agonists and Partial Agonists. J. Biol. Chem. 2005, 280, 22165−22171. (105) Schertler, G. F. X.; Villa, C.; Henderson, R. Projection Structure of Rhodopsin. Nature 1993, 362, 770−772. (106) Palczewski, K.; Kumasaka, T.; Hori, T.; Behnke, C. A.; Motoshima, H.; Fox, B. A.; Le Trong, I.; Teller, D. C.; Okada, T.; Stenkamp, R. E.; et al. Crystal Structure of Rhodopsin: A G ProteinCoupled Receptor. Science 2000, 289, 739−745. (107) Teller, D. C.; Okada, T.; Behnke, C. A.; Palczewski, K.; Stenkamp, R. E. Advances in Determination of a High-Resolution Three-Dimensional Structure of Rhodopsin, a Model of G-ProteinCoupled Receptors (GPCRs). Biochemistry 2001, 40, 7761−7772. (108) Li, J.; Edwards, P. C.; Burghammer, M.; Villa, C.; Schertler, G. F. Structure of Bovine Rhodopsin in a Trigonal Crystal Form. J. Mol. Biol. 2004, 343, 1409−1438. (109) Okada, T.; Sugihara, M.; Bondar, A. N.; Elstner, M.; Entel, P.; Buss, V. The Retinal Conformation and Its Environment in Rhodopsin in Light of a New 2.2 Å Crystal Structure. J. Mol. Biol. 2004, 342, 571− 583. (110) Park, J. H.; Scheerer, P.; Hofmann, K. P.; Choe, H. W.; Ernst, O. P. Crystal Structure of the Ligand-Free G-Protein-Coupled Receptor Opsin. Nature 2008, 454, 183−187. (111) Scheerer, P.; Park, J. H.; Hildebrand, P. W.; Kim, Y. J.; Krauss, N.; Choe, H. W.; Hofmann, K. P.; Ernst, O. P. Crystal Structure of Opsin in Its G-Protein-Interacting Conformation. Nature 2008, 455, 497−502.

(112) Choe, H. W.; Kim, Y. J.; Park, J. H.; Morizumi, T.; Pai, E. F.; Krauss, N.; Hofmann, K. P.; Scheerer, P.; Ernst, O. P. Crystal Structure of Metarhodopsin II. Nature 2011, 471, 651−655. (113) Standfuss, J.; Edwards, P. C.; D’Antona, A.; Fransen, M.; Xie, G.; Oprian, D. D.; Schertler, G. F. The Structural Basis of AgonistInduced Activation in Constitutively Active Rhodopsin. Nature 2011, 471, 656−660. (114) Kang, Y.; Zhou, X. E.; Gao, X.; He, Y.; Liu, W.; Ishchenko, A.; Barty, A.; White, T. A.; Yefanov, O.; Han, G. W.; et al. Crystal Structure of Rhodopsin Bound to Arrestin by Femtosecond X-Ray Laser. Nature 2015, 523, 561−567. (115) Rosenbaum, D. M.; Cherezov, V.; Hanson, M. A.; Rasmussen, S. G.; Thian, F. S.; Kobilka, T. S.; Choi, H. J.; Yao, X. J.; Weis, W. I.; Stevens, R. C.; et al. GPCR Engineering Yields High-Resolution Structural Insights into β(2)-Adrenergic Receptor Function. Science 2007, 318, 1266−1273. (116) Rasmussen, S. G.; Choi, H. J.; Rosenbaum, D. M.; Kobilka, T. S.; Thian, F. S.; Edwards, P. C.; Burghammer, M.; Ratnala, V. R.; Sanishvili, R.; Fischetti, R. F.; et al. Crystal Structure of the Human β(2) Adrenergic G-Protein-Coupled Receptor. Nature 2007, 450, 383−387. (117) Cherezov, V.; Rosenbaum, D. M.; Hanson, M. A.; Rasmussen, S. G.; Thian, F. S.; Kobilka, T. S.; Choi, H. J.; Kuhn, P.; Weis, W. I.; Kobilka, B. K.; et al. High-Resolution Crystal Structure of an Engineered Human β2-Adrenergic G Protein-Coupled Receptor. Science 2007, 318, 1258−1265. (118) Rasmussen, S. G.; Choi, H. J.; Fung, J. J.; Pardon, E.; Casarosa, P.; Chae, P. S.; Devree, B. T.; Rosenbaum, D. M.; Thian, F. S.; Kobilka, T. S.; et al. Structure of a Nanobody-Stabilized Active State of the β(2) Adrenoceptor. Nature 2011, 469, 175−180. (119) Rasmussen, S. G.; DeVree, B. T.; Zou, Y.; Kruse, A. C.; Chung, K. Y.; Kobilka, T. S.; Thian, F. S.; Chae, P. S.; Pardon, E.; Calinski, D.; et al. Crystal Structure of the β(2) Adrenergic Receptor-Gs Protein Complex. Nature 2011, 477, 549−555. (120) Deupi, X.; Standfuss, J.; Schertler, G. Conserved Activation Pathways in G Protein-Coupled Receptors. Biochem. Soc. Trans. 2012, 40, 383−388. (121) Deupi, X. Relevance of Rhodopsin Studies for GPCR Activation. Biochim. Biophys. Acta, Bioenerg. 2014, 1837, 674−682. (122) Jaakola, V. P.; Griffith, M. T.; Hanson, M. A.; Cherezov, V.; Chien, E. Y.; Lane, J. R.; Ijzerman, A. P.; Stevens, R. C. The 2.6 Angstrom Crystal Structure of a Human A2a Adenosine Receptor Bound to an Antagonist. Science 2008, 322, 1211−1217. (123) Lebon, G.; Warne, T.; Edwards, P. C.; Bennett, K.; Langmead, C. J.; Leslie, A. G.; Tate, C. G. Agonist-Bound Adenosine A2a Receptor Structures Reveal Common Features of Gpcr Activation. Nature 2011, 474, 521−525. (124) Xu, F.; Wu, H.; Katritch, V.; Han, G. W.; Jacobson, K. A.; Gao, Z. G.; Cherezov, V.; Stevens, R. C. Structure of an Agonist-Bound Human A2a Adenosine Receptor. Science 2011, 332, 322−327. (125) Kruse, A. C.; Hu, J.; Pan, A. C.; Arlow, D. H.; Rosenbaum, D. M.; Rosemond, E.; Green, H. F.; Liu, T.; Chae, P. S.; Dror, R. O.; et al. Structure and Dynamics of the M3Muscarinic Acetylcholine Receptor. Nature 2012, 482, 552−556. (126) Kruse, A. C.; Ring, A. M.; Manglik, A.; Hu, J.; Hu, K.; Eitel, K.; Hubner, H.; Pardon, E.; Valant, C.; Sexton, P. M.; et al. Activation and Allosteric Modulation of a Muscarinic Acetylcholine Receptor. Nature 2013, 504, 101−106. (127) Manglik, A.; Kruse, A. C.; Kobilka, T. S.; Thian, F. S.; Mathiesen, J. M.; Sunahara, R. K.; Pardo, L.; Weis, W. I.; Kobilka, B. K.; Granier, S. Crystal Structure of the μ-Opioid Receptor Bound to a Morphinan Antagonist. Nature 2012, 485, 321−326. (128) Huang, W.; Manglik, A.; Venkatakrishnan, A. J.; Laeremans, T.; Feinberg, E. N.; Sanborn, A. L.; Kato, H. E.; Livingston, K. E.; Thorsen, T. S.; Kling, R. C.; et al. Structural Insights into μ-Opioid Receptor Activation. Nature 2015, 524, 315−321. (129) Katritch, V.; Cherezov, V.; Stevens, R. C. Diversity and Modularity of G Protein-Coupled Receptor Structures. Trends Pharmacol. Sci. 2012, 33, 17−27. AS

DOI: 10.1021/acs.chemrev.6b00084 Chem. Rev. XXXX, XXX, XXX−XXX

Chemical Reviews

Review

(130) Katritch, V.; Cherezov, V.; Stevens, R. C. Structure-Function of the G Protein-Coupled Receptor Superfamily. Annu. Rev. Pharmacol. Toxicol. 2013, 53, 531−556. (131) Venkatakrishnan, A. J.; Deupi, X.; Lebon, G.; Tate, C. G.; Schertler, G. F.; Babu, M. M. Molecular Signatures of G ProteinCoupled Receptors. Nature 2013, 494, 185−194. (132) Frauenfelder, H.; Parak, F.; Young, R. D. Conformational Substates in Proteins. Annu. Rev. Biophys. Biophys. Chem. 1988, 17, 451−479. (133) Frauenfelder, H.; Sligar, S. G.; Wolynes, P. G. The Energy Landscapes and Motions of Proteins. Science 1991, 254, 1598−1603. (134) Henzler-Wildman, K.; Kern, D. Dynamic Personalities of Proteins. Nature 2007, 450, 964−972. (135) Kobilka, B. K.; Deupi, X. Conformational Complexity of G Protein-Coupled Receptors. Trends Pharmacol. Sci. 2007, 28, 397−406. (136) Deupi, X.; Kobilka, B. K. Energy Landscapes as a Tool to Integrate GPCR Structure, Dynamics, and Function. Physiology 2010, 25, 293−303. (137) Rosenbaum, D. M.; Rasmussen, S. G. F.; Kobilka, B. K. The Structure and Function of G Protein-Coupled Receptors. Nature 2009, 459, 356−363. (138) Vaidehi, N.; Kenakin, T. The Role of Conformational Ensembles of Seven Transmembrane Receptors in Functional Selectivity. Curr. Opin. Pharmacol. 2010, 10, 775−781. (139) Manglik, A.; Kobilka, B. The Role of Protein Dynamics in GPCR Function: Insights from the β(2)AR and Rhodopsin. Curr. Opin. Cell Biol. 2014, 27, 136−143. (140) Reiter, E.; Ahn, S.; Shukla, A. K.; Lefkowitz, R. J. Molecular Mechanism of β-Arrestin-Biased Agonism at Seven-Transmembrane Receptors. Annu. Rev. Pharmacol. Toxicol. 2012, 52, 179−197. (141) Kenakin, T.; Christopoulos, A. Signalling Bias in New Drug Discovery: Detection, Quantification and Therapeutic Impact. Nat. Rev. Drug Discovery 2012, 12, 205−216. (142) Wisler, J. W.; Xiao, K.; Thomsen, A. R.; Lefkowitz, R. J. Recent Developments in Biased Agonism. Curr. Opin. Cell Biol. 2014, 27, 18− 24. (143) Heldin, C. H. Dimerization of Cell-Surface Receptors in Signal-Transduction. Cell 1995, 80, 213−223. (144) Whorton, M. R.; Bokoch, M. P.; Rasmussen, S. G. F.; Huang, B.; Zare, R. N.; Kobilka, B.; Sunahara, R. K. A Monomeric G ProteinCoupled Receptor Isolated in a High-Density Lipoprotein Particle Efficiently Activates Its G Protein. Proc. Natl. Acad. Sci. U. S. A. 2007, 104, 7682−7687. (145) Banerjee, S.; Huber, T.; Sakmar, T. P. Rapid Incorporation of Functional Rhodopsin into Nanoscale Apolipoprotein Bound Bilayer (NABB) Particles. J. Mol. Biol. 2008, 377, 1067−1081. (146) Whorton, M. R.; Jastrzebska, B.; Park, P. S.; Fotiadis, D.; Engel, A.; Palczewski, K.; Sunahara, R. K. Efficient Coupling of Transducin to Monomeric Rhodopsin in a Phospholipid Bilayer. J. Biol. Chem. 2008, 283, 4387−4394. (147) Ernst, O. P.; Gramse, V.; Kolbe, M.; Hofmann, K. P.; Heck, M. Monomeric G Protein-Coupled Receptor Rhodopsin in Solution Activates Its G Protein Transducin at the Diffusion Limit. Proc. Natl. Acad. Sci. U. S. A. 2007, 104, 10859−10864. (148) Bouvier, M. Oligomerization of G Protein-Coupled Transmitter Receptors. Nat. Rev. Neurosci. 2001, 2, 274−286. (149) Angers, S.; Salahpour, A.; Bouvier, M. Dimerization: An Emerging Concept for G Protein-Coupled Receptor Ontogeny and Function. Annu. Rev. Pharmacol. Toxicol. 2002, 42, 409−435. (150) Terrillon, S.; Bouvier, M. Roles of G Protein-Coupled Receptor Dimerization - from Ontogeny to Signalling Regulation. EMBO Rep. 2004, 5, 30−34. (151) Lohse, M. J. Dimerization in GPCR Mobility and Signaling. Curr. Opin. Pharmacol. 2010, 10, 53−58. (152) Botelho, A. V.; Huber, T.; Sakmar, T. P.; Brown, M. F. Curvature and Hydrophobic Forces Drive Oligomerization and Modulate Activity of Rhodopsin in Membranes. Biophys. J. 2006, 91, 4464−4477.

(153) Mansoor, S. E.; Palczewski, K.; Farrens, D. L. Rhodopsin SelfAssociates in Asolectin Liposomes. Proc. Natl. Acad. Sci. U. S. A. 2006, 103, 3060−3065. (154) Fung, J. J.; Deupi, X.; Pardo, L.; Yao, X. J.; Velez-Ruiz, G. A.; DeVree, B. T.; Sunahara, R. K.; Kobilka, B. K. Ligand-Regulated Oligomerization of β(2)-Adrenoceptors in a Model Lipid Bilayer. EMBO J. 2009, 28, 3315−3328. (155) Fotiadis, D.; Liang, Y.; Filipek, S.; Saperstein, D. A.; Engel, A.; Palczewski, K. Atomic-Force Microscopy: Rhodopsin Dimers in Native Disc Membranes. Nature 2003, 421, 127−128. (156) Liang, Y.; Fotiadis, D.; Filipek, S.; Saperstein, D. A.; Palczewski, K.; Engel, A. Organization of the G Protein-Coupled Receptors Rhodopsin and Opsin in Native Membranes. J. Biol. Chem. 2003, 278, 21655−21662. (157) Ianoul, A.; Grant, D. D.; Rouleau, Y.; Bani-Yaghoub, M.; Johnston, L. J.; Pezacki, J. P. Imaging Nanometer Domains of βAdrenergic Receptor Complexes on the Surface of Cardiac Myocytes. Nat. Chem. Biol. 2005, 1, 196−202. (158) Dorsch, S.; Klotz, K. N.; Engelhardt, S.; Lohse, M. J.; Bunemann, M. Analysis of Receptor Oligomerization by FRAP Microscopy. Nat. Methods 2009, 6, 225−230. (159) Albizu, L.; Cottet, M.; Kralikova, M.; Stoev, S.; Seyer, R.; Brabet, I.; Roux, T.; Bazin, H.; Bourrier, E.; Lamarque, L.; et al. TimeResolved FRET between GPCR Ligands Reveals Oligomers in Native Tissues. Nat. Chem. Biol. 2010, 6, 587−594. (160) Gurevich, V. V.; Gurevich, E. V. GPCR Monomers and Oligomers: It Takes All Kinds. Trends Neurosci. 2008, 31, 74−81. (161) Ferre, S.; Casado, V.; Devi, L. A.; Filizola, M.; Jockers, R.; Lohse, M. J.; Milligan, G.; Pin, J. P.; Guitart, X. G Protein-Coupled Receptor Oligomerization Revisited: Functional and Pharmacological Perspectives. Pharmacol. Rev. 2014, 66, 413−434. (162) Mathiasen, S.; Tonnesen, A.; Christensen, S.; Fung, J. J.; Rasmussen, S. G. F.; Borrero, E.; Provasi, D.; Filizola, M.; Kobilka, B.; Stamou, D. Membrane Curvature Regulates the Oligomerization of Human β(2)-Adrenergic Receptors. Biophys. J. 2013, 104, 42A−42A. (163) Margeta-Mitrovic, M.; Jan, Y. N.; Jan, L. Y. A Trafficking Checkpoint Controls GABA(B) Receptor Heterodimerization. Neuron 2000, 27, 97−106. (164) Salahpour, A.; Angers, S.; Mercier, J. F.; Lagace, M.; Marullo, S.; Bouvier, M. Homodimerization of the β(2)-Adrenergic Receptor as a Prerequisite for Cell Surface Targeting. J. Biol. Chem. 2004, 279, 33390−33397. (165) Hague, C.; Uberti, M. A.; Chen, Z. J.; Hall, R. A.; Minneman, K. P. Cell Surface Expression of α(1d)-Adrenergic Receptors Is Controlled by Heterodimerization with α(1b)-Adrenergic Receptors. J. Biol. Chem. 2004, 279, 15541−15549. (166) Maurice, P.; Kamal, M.; Jockers, R. Asymmetry of GPCR Oligomers Supports Their Functional Relevance. Trends Pharmacol. Sci. 2011, 32, 514−520. (167) George, S. R.; O’Dowd, B. F.; Lee, S. R. G Protein-Coupled Receptor Oligomerization and Its Potential for Drug Discovery. Nat. Rev. Drug Discovery 2002, 1, 808−820. (168) Shaner, N. C.; Steinbach, P. A.; Tsien, R. Y. A Guide to Choosing Fluorescent Proteins. Nat. Methods 2005, 2, 905−909. (169) Goncalves, M. S. Fluorescent Labeling of Biomolecules with Organic Probes. Chem. Rev. 2009, 109, 190−212. (170) Selvin, P. R. Principles and Biophysical Applications of Lanthanide-Based Probes. Annu. Rev. Biophys. Biomol. Struct. 2002, 31, 275−302. (171) Alivisatos, A. P.; Gu, W. W.; Larabell, C. Quantum Dots as Cellular Probes. Annu. Rev. Biomed. Eng. 2005, 7, 55−76. (172) Kobilka, B. K.; Kobilka, T. S.; Daniel, K.; Regan, J. W.; Caron, M. G.; Lefkowitz, R. J. Chimeric α(2)-, β(2)-Adrenergic Receptors: Delineation of Domains Involved in Effector Coupling and Ligand Binding Specificity. Science 1988, 240, 1310−1316. (173) Prescher, J. A.; Bertozzi, C. R. Chemistry in Living Systems. Nat. Chem. Biol. 2005, 1, 13−21. AT

DOI: 10.1021/acs.chemrev.6b00084 Chem. Rev. XXXX, XXX, XXX−XXX

Chemical Reviews

Review

(174) Sletten, E. M.; Bertozzi, C. R. Bioorthogonal Chemistry: Fishing for Selectivity in a Sea of Functionality. Angew. Chem., Int. Ed. 2009, 48, 6974−6998. (175) Tan, C. M.; Brady, A. E.; Nickols, H. H.; Wang, Q.; Limbird, L. E. Membrane Trafficking of G Protein-Coupled Receptors. Annu. Rev. Pharmacol. Toxicol. 2004, 44, 559−609. (176) Coons, A. H. The Beginnings of Immunofluorescence. J. Immunol. 1961, 87, 499−503. (177) Fraser, C. M.; Venter, J. C. Monoclonal Antibodies to βAdrenergic Receptors: Use in Purification and Molecular Characterization of β Receptors. Proc. Natl. Acad. Sci. U. S. A. 1980, 77, 7034− 7038. (178) Molday, R. S.; MacKenzie, D. Monoclonal Antibodies to Rhodopsin: Characterization, Cross-Reactivity, and Application as Structural Probes. Biochemistry 1983, 22, 653−660. (179) Couraud, P.-O.; Delavier-Klutchko, C.; Durieu-Trautmann, O.; Strosberg, A. D. Antibodies Raised against β-Adrenergic Receptors Stimulate Adenylate Cyclase. Biochem. Biophys. Res. Commun. 1981, 99, 1295−1302. (180) Blanpain, C.; Vanderwinden, J. M.; Cihak, J.; Wittamer, V.; Le Poul, E.; Issafras, H.; Stangassinger, M.; Vassart, G.; Marullo, S.; Schlondorff, D.; et al. Multiple Active States and Oligomerization of CCR5 Revealed by Functional Properties of Monoclonal Antibodies. Mol. Biol. Cell 2002, 13, 723−737. (181) Issafras, H.; Angers, S.; Bulenger, S.; Blanpain, C.; Parmentier, M.; Labbe-Jullie, C.; Bouvier, M.; Marullo, S. Constitutive AgonistIndependent CCR5 Oligomerization and Antibody-Mediated Clustering Occurring at Physiological Levels of Receptors. J. Biol. Chem. 2002, 277, 34666−34673. (182) Munro, S.; Pelham, H. R. Use of Peptide Tagging to Detect Proteins Expressed from Cloned Genes: Deletion Mapping Functional Domains of Drosophila Hsp 70. EMBO J. 1984, 3, 3087−3093. (183) Young, C. L.; Britton, Z. T.; Robinson, A. S. Recombinant Protein Expression and Purification: A Comprehensive Review of Affinity Tags and Microbial Applications. Biotechnol. J. 2012, 7, 620− 634. (184) Lameh, J.; Philip, M.; Sharma, Y. K.; Moro, O.; Ramachandran, J.; Sadee, W. Hm1Muscarinic Cholinergic Receptor Internalization Requires a Domain in the 3rd Cytoplasmic Loop. J. Biol. Chem. 1992, 267, 13406−13412. (185) Hadcock, J. R.; Wang, H. Y.; Malbon, C. C. Agonist-Induced Destabilization of β-Adrenergic-Receptor Messenger-RNA - Attenuation of Glucocorticoid-Induced up-Regulation of β-Adrenergic Receptors. J. Biol. Chem. 1989, 264, 19928−19933. (186) Zhang, J.; Ferguson, S. G.; Barak, L. S.; Menard, L.; Caron, M. G. Dynamin and β-Arrestin Reveal Distinct Mechanisms for G Protein-Coupled Receptor Internalization. J. Biol. Chem. 1996, 271, 18302−18305. (187) Rocheville, M.; Lange, D. C.; Kumar, U.; Sasi, R.; Patel, R. C.; Patel, Y. C. Subtypes of the Somatostatin Receptor Assemble as Functional Homo- and Heterodimers. J. Biol. Chem. 2000, 275, 7862− 7869. (188) Rocheville, M.; Lange, D. C.; Kumar, U.; Patel, S. C.; Patel, R. C.; Patel, Y. C. Receptors for Dopamine and Somatostatin: Formation of Hetero-Oligomers with Enhanced Functional Activity. Science 2000, 288, 154−157. (189) McVey, M.; Ramsay, D.; Kellett, E.; Rees, S.; Wilson, S.; Pope, A. J.; Milligan, G. Monitoring Receptor Oligomerization Using TimeResolved Fluorescence Resonance Energy Transfer and Bioluminescence Resonance Energy Transfer - the Human δ-Opioid Receptor Displays Constitutive Oligomerization at the Cell Surface, Which Is Not Regulated by Receptor Occupancy. J. Biol. Chem. 2001, 276, 14092−14099. (190) Wu, L.; LaRosa, G.; Kassam, N.; Gordon, C. J.; Heath, H.; Ruffing, N.; Chen, H.; Humblias, J.; Samson, M.; Parmentier, M.; et al. Interaction of Chemokine Receptor CCR5 with Its Ligands: Multiple Domains for HIV-1 gp120 Binding and a Single Domain for Chemokine Binding. J. Exp. Med. 1997, 186, 1373−1381.

(191) Gupta, A.; Decaillot, F. M.; Gomes, I.; Tkalych, O.; Heimann, A. S.; Ferro, E. S.; Devi, L. A. Conformation State-Sensitive Antibodies to G Protein-Coupled Receptors. J. Biol. Chem. 2007, 282, 5116−5124. (192) Mancia, F.; Brenner-Morton, S.; Siegel, R.; Assur, Z.; Sun, Y.; Schieren, I.; Mendelsohn, M.; Axel, R.; Hendrickson, W. A. Production and Characterization of Monoclonal Antibodies Sensitive to Conformation in the 5HT2c Serotonin Receptor. Proc. Natl. Acad. Sci. U. S. A. 2007, 104, 4303−4308. (193) Hutchings, C. J.; Koglin, M.; Marshall, F. H. Therapeutic Antibodies Directed at G Protein-Coupled Receptors. mAbs 2010, 2, 594−606. (194) Navratilova, I.; Sodroski, J.; Myszka, D. G. Solubilization, Stabilization, and Purification of Chemokine Receptors Using Biosensor Technology. Anal. Biochem. 2005, 339, 271−281. (195) Steyaert, J.; Kobilka, B. K. Nanobody Stabilization of G Protein-Coupled Receptor Conformational States. Curr. Opin. Struct. Biol. 2011, 21, 567−572. (196) Ring, A. M.; Manglik, A.; Kruse, A. C.; Enos, M. D.; Weis, W. I.; Garcia, K. C.; Kobilka, B. K. Adrenaline-Activated Structure of β2Adrenoceptor Stabilized by an Engineered Nanobody. Nature 2013, 502, 575−579. (197) Vaneycken, I.; D’huyvetter, M.; Hernot, S.; De Vos, J.; Xavier, C.; Devoogdt, N.; Caveliers, V.; Lahoutte, T. Immuno-Imaging Using Nanobodies. Curr. Opin. Biotechnol. 2011, 22, 877−881. (198) Irannejad, R.; Tomshine, J. C.; Tomshine, J. R.; Chevalier, M.; Mahoney, J. P.; Steyaert, J.; Rasmussen, S. G.; Sunahara, R. K.; ElSamad, H.; Huang, B.; et al. Conformational Biosensors Reveal GPCR Signalling from Endosomes. Nature 2013, 495, 534−538. (199) Lefkowitz, R. J.; Mukherjee, C.; Coverstone, M.; Caron, M. G. Stereospecific (3H)(−)-Alprenolol Binding Sites, β-Adrenergic Receptors and Adenylate Cyclase. Biochem. Biophys. Res. Commun. 1974, 60, 703−709. (200) Williams, L. T.; Lefkowitz, R. J. α-Adrenergic Receptor Identification by (3H)Dihydroergocryptine Binding. Science 1976, 192, 791−793. (201) Knepp, A. M.; Grunbeck, A.; Banerjee, S.; Sakmar, T. P.; Huber, T. Direct Measurement of Thermal Stability of Expressed CCR5 and Stabilization by Small Molecule Ligands. Biochemistry 2011, 50, 502−511. (202) Middleton, R. J.; Kellam, B. Fluorophore-Tagged GPCR Ligands. Curr. Opin. Chem. Biol. 2005, 9, 517−525. (203) Daly, C. J.; Ross, R. A.; Whyte, J.; Henstridge, C. M.; Irving, A. J.; McGrath, J. C. Fluorescent Ligand Binding Reveals Heterogeneous Distribution of Adrenoceptors and ’Cannabinoid-Like’ Receptors in Small Arteries. Br. J. Pharmacol. 2010, 159, 787−796. (204) Sridharan, R.; Zuber, J.; Connelly, S. M.; Mathew, E.; Dumont, M. E. Fluorescent Approaches for Understanding Interactions of Ligands with G Protein-Coupled Receptors. Biochim. Biophys. Acta, Biomembr. 2014, 1838, 15−33. (205) Ilien, B.; Franchet, C.; Bernard, P.; Morisset, S.; Weill, C. O.; Bourguignon, J. J.; Hibert, M.; Galzi, J. L. Fluorescence Resonance Energy Transfer to Probe Human M1Muscarinic Receptor Structure and Drug Binding Properties. J. Neurochem. 2003, 85, 768−778. (206) Huber, T.; Sakmar, T. P. Chemical Biology Methods for Investigating G Protein-Coupled Receptor Signaling. Chem. Biol. 2014, 21, 1224−1237. (207) James, J. R.; Oliveira, M. I.; Carmo, A. M.; Iaboni, A.; Davis, S. J. A Rigorous Experimental Framework for Detecting Protein Oligomerization Using Bioluminescence Resonance Energy Transfer. Nat. Methods 2006, 3, 1001−1006. (208) Bouvier, M.; Heveker, N.; Jockers, R.; Marullo, S.; Milligan, G. BRET Analysis of GPCR Oligomerization: Newer Does Not Mean Better. Nat. Methods 2007, 4, 3−4. (209) Cottet, M.; Albizu, L.; Comps-Agrar, L.; Trinquet, E.; Pin, J. P.; Mouillac, B.; Durroux, T. A Rigorous Experimental Framework for Detecting Protein Oligomerization Using Bioluminescence Resonance Energy Transfer. Methods Mol. Biol. 2011, 746, 373−387. (210) Zwier, J. M.; Roux, T.; Cottet, M.; Durroux, T.; Douzon, S.; Bdioui, S.; Gregor, N.; Bourrier, E.; Oueslati, N.; Nicolas, L.; et al. A AU

DOI: 10.1021/acs.chemrev.6b00084 Chem. Rev. XXXX, XXX, XXX−XXX

Chemical Reviews

Review

Functionally Intact β(2)-Adrenergic Receptor-Green Fluorescent Protein Conjugate. Mol. Pharmacol. 1997, 51, 177−184. (231) Vilardaga, J. P.; Bunemann, M.; Krasel, C.; Castro, M.; Lohse, M. J. Measurement of the Millisecond Activation Switch of G ProteinCoupled Receptors in Living Cells. Nat. Biotechnol. 2003, 21, 807− 812. (232) Milligan, G. Exploring the Dynamics of Regulation of G Protein-Coupled Receptors Using Green Fluorescent Protein. Br. J. Pharmacol. 1999, 128, 501−510. (233) Kallal, L.; Benovic, J. L. Using Green Fluorescent Proteins to Study G Protein-Coupled Receptor Localization and Trafficking. Trends Pharmacol. Sci. 2000, 21, 175−180. (234) Gether, U.; Lin, S.; Kobilka, B. K. Fluorescent Labeling of Purified β(2) Adrenergic Receptor: Evidence for Ligand-Specific Conformational Changes. J. Biol. Chem. 1995, 270, 28268−28275. (235) Overton, M. C.; Blumer, K. J. G Protein-Coupled Receptors Function as Oligomers in Vivo. Curr. Biol. 2000, 10, 341−344. (236) Tarasova, N. I.; Stauber, R. H.; Choi, J. K.; Hudson, E. A.; Czerwinski, G.; Miller, J. L.; Pavlakis, G. N.; Michejda, C. J.; Wank, S. A. Visualization of G Protein-Coupled Receptor Trafficking with the Aid of the Green Fluorescent Protein. Endocytosis and Recycling of Cholecystokinin Receptor Type A. J. Biol. Chem. 1997, 272, 14817− 14824. (237) Xiao, Z.; Zhang, N.; Murphy, D. B.; Devreotes, P. N. Dynamic Distribution of Chemoattractant Receptors in Living Cells During Chemotaxis and Persistent Stimulation. J. Cell Biol. 1997, 139, 365− 374. (238) McLean, A. J.; Milligan, G. Ligand Regulation of Green Fluorescent Protein-Tagged Forms of the Human β(1)- and β(2)Adrenoceptors; Comparisons with the Unmodified Receptors. Br. J. Pharmacol. 2000, 130, 1825−1832. (239) Greasley, P. J.; Fanelli, F.; Scheer, A.; Abuin, L.; NennigerTosato, M.; DeBenedetti, P. G.; Cotecchia, S. Mutational and Computational Analysis of the α(1b)-Adrenergic Receptor: Involvement of Basic and Hydrophobic Residues in Receptor Activation and G Protein Coupling. J. Biol. Chem. 2001, 276, 46485−46494. (240) Baird, G. S.; Zacharias, D. A.; Tsien, R. Y. Circular Permutation and Receptor Insertion within Green Fluorescent Proteins. Proc. Natl. Acad. Sci. U. S. A. 1999, 96, 11241−11246. (241) Cabantous, S.; Terwilliger, T. C.; Waldo, G. S. Protein Tagging and Detection with Engineered Self-Assembling Fragments of Green Fluorescent Protein. Nat. Biotechnol. 2005, 23, 102−107. (242) Jiang, W. X.; Dong, X.; Jiang, J.; Yang, Y. H.; Yang, J.; Lu, Y. B.; Fang, S. H.; Wei, E. Q.; Tang, C.; Zhang, W. P. Specific Cell Surface Labeling of GPCRs Using Split GFP. Sci. Rep. 2016, 6, 20568. (243) Pfleger, K. D. G.; Eidne, K. A. Illuminating Insights into Protein-Protein Interactions Using Bioluminescence Resonance Energy Transfer (BRET). Nat. Methods 2006, 3, 165−174. (244) Prinz, A.; Diskar, M.; Herberg, F. W. Application of Bioluminescence Resonance Energy Transfer (BRET) for Biomolecular Interaction Studies. ChemBioChem 2006, 7, 1007−1012. (245) Angers, S.; Salahpour, A.; Joly, E.; Hilairet, S.; Chelsky, D.; Dennis, M.; Bouvier, M. Detection of β(2)-Adrenergic Receptor Dimerization in Living Cells Using Bioluminescence Resonance Energy Transfer (BRET). Proc. Natl. Acad. Sci. U. S. A. 2000, 97, 3684−3689. (246) Kroeger, K. M.; Hanyaloglu, A. C.; Seeber, R. M.; Miles, L. E.; Eidne, K. A. Constitutive and Agonist-Dependent Homo-Oligomerization of the Thyrotropin-Releasing Hormone Receptor: Detection in Living Cells Using Bioluminescence Resonance Energy Transfer. J. Biol. Chem. 2001, 276, 12736−12743. (247) Mercier, J. F.; Salahpour, A.; Angers, S.; Breit, A.; Bouvier, M. Quantitative Assessment of β(1)- and β(2)-Adrenergic Receptor Homo- and Heterodimerization by Bioluminescence Resonance Energy Transfer. J. Biol. Chem. 2002, 277, 44925−44931. (248) Ciruela, F.; Fernandez-Duenas, V. GPCR Oligomerization Analysis by Means of BRET and dFRAP. Methods Mol. Biol. 2015, 1272, 133−141.

Fluorescent Ligand-Binding Alternative Using Tag-Lite (R) Technology. J. Biomol. Screening 2010, 15, 1248−1259. (211) Emami-Nemini, A.; Roux, T.; Leblay, M.; Bourrier, E.; Lamarque, L.; Trinquet, E.; Lohse, M. J. Time-Resolved Fluorescence Ligand Binding for G Protein-Coupled Receptors. Nat. Protoc. 2013, 8, 1307−1320. (212) Leyris, J. P.; Roux, T.; Trinquet, E.; Verdie, P.; Fehrentz, J. A.; Oueslati, N.; Douzon, S.; Bourrier, E.; Lamarque, L.; Gagne, D.; et al. Homogeneous Time-Resolved Fluorescence-Based Assay to Screen for Ligands Targeting the Growth Hormone Secretagogue Receptor Type 1a. Anal. Biochem. 2011, 408, 253−262. (213) Tan, Q.; Zhu, Y.; Li, J.; Chen, Z.; Han, G. W.; Kufareva, I.; Li, T.; Ma, L.; Fenalti, G.; Zhang, W.; et al. Structure of the CCR5 Chemokine Receptor-Hiv Entry Inhibitor Maraviroc Complex. Science 2013, 341, 1387−1390. (214) Shonberg, J.; Scammells, P. J.; Capuano, B. Design Strategies for Bivalent Ligands Targeting GPCRs. ChemMedChem 2011, 6, 963− 974. (215) Tanaka, T.; Nomura, W.; Narumi, T.; Masuda, A.; Tamamura, H. Bivalent Ligands of CXCR4 with Rigid Linkers for Elucidation of the Dimerization State in Cells. J. Am. Chem. Soc. 2010, 132, 15899− 15901. (216) Kuder, K.; Kiec-Kononowicz, K. Fluorescent GPCR Ligands as New Tools in Pharmacology. Curr. Med. Chem. 2008, 15, 2132−2143. (217) Böhme, I.; Beck-Sickinger, A. G. Illuminating the Life of GPCRs. Cell Commun. Signaling 2009, 7, 16. (218) Kuder, K. J.; Kiec-Kononowicz, K. Fluorescent GPCR Ligands as New Tools in Pharmacology-Update, Years 2008-Early 2014. Curr. Med. Chem. 2014, 21, 3962−3975. (219) Hayashi, T.; Hamachi, I. Traceless Affinity Labeling of Endogenous Proteins for Functional Analysis in Living Cells. Acc. Chem. Res. 2012, 45, 1460−1469. (220) Dohlman, H. G.; Caron, M. G.; Strader, C. D.; Amlaiky, N.; Lefkowitz, R. J. Identification and Sequence of a Binding-Site Peptide of the β(2) Adrenergic Receptor. Biochemistry 1988, 27, 1813−1817. (221) Rosenbaum, D. M.; Zhang, C.; Lyons, J. A.; Holl, R.; Aragao, D.; Arlow, D. H.; Rasmussen, S. G.; Choi, H. J.; Devree, B. T.; Sunahara, R. K.; et al. Structure and Function of an Irreversible Agonist-β(2) Adrenoceptor Complex. Nature 2011, 469, 236−240. (222) Weichert, D.; Kruse, A. C.; Manglik, A.; Hiller, C.; Zhang, C.; Hubner, H.; Kobilka, B. K.; Gmeiner, P. Covalent Agonists for Studying G Protein-Coupled Receptor Activation. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, 10744−10748. (223) Tsukiji, S.; Miyagawa, M.; Takaoka, Y.; Tamura, T.; Hamachi, I. Ligand-Directed Tosyl Chemistry for Protein Labeling in Vivo. Nat. Chem. Biol. 2009, 5, 341−343. (224) Fujishima, S. H.; Yasui, R.; Miki, T.; Ojida, A.; Hamachi, I. Ligand-Directed Acyl Imidazole Chemistry for Labeling of MembraneBound Proteins on Live Cells. J. Am. Chem. Soc. 2012, 134, 3961− 3964. (225) Miki, T.; Fujishima, S.; Komatsu, K.; Kuwata, K.; Kiyonaka, S.; Hamachi, I. LDAI-Based Chemical Labeling of Intact Membrane Proteins and Its Pulse-Chase Analysis under Live Cell Conditions. Chem. Biol. 2014, 21, 1013−1022. (226) Chalfie, M.; Tu, Y.; Euskirchen, G.; Ward, W. W.; Prasher, D. C. Green Fluorescent Protein as a Marker for Gene Expression. Science 1994, 263, 802−805. (227) Heim, R.; Prasher, D. C.; Tsien, R. Y. Wavelength Mutations and Posttranslational Autoxidation of Green Fluorescent Protein. Proc. Natl. Acad. Sci. U. S. A. 1994, 91, 12501−12504. (228) Cubitt, A. B.; Heim, R.; Adams, S. R.; Boyd, A. E.; Gross, L. A.; Tsien, R. Y. Understanding, Improving and Using Green Fluorescent Proteins. Trends Biochem. Sci. 1995, 20, 448−455. (229) Giepmans, B. N.; Adams, S. R.; Ellisman, M. H.; Tsien, R. Y. The Fluorescent Toolbox for Assessing Protein Location and Function. Science 2006, 312, 217−224. (230) Barak, L. S.; Ferguson, S. S.; Zhang, J.; Martenson, C.; Meyer, T.; Caron, M. G. Internal Trafficking and Surface Mobility of a AV

DOI: 10.1021/acs.chemrev.6b00084 Chem. Rev. XXXX, XXX, XXX−XXX

Chemical Reviews

Review

(249) Milligan, G.; Bouvier, M. Methods to Monitor the Quaternary Structure of G Protein-Coupled Receptors. FEBS J. 2005, 272, 2914− 2925. (250) Lohse, M. J.; Nuber, S.; Hoffmann, C. Fluorescence/ Bioluminescence Resonance Energy Transfer Techniques to Study G Protein-Coupled Receptor Activation and Signaling. Pharmacol. Rev. 2012, 64, 299−336. (251) Kocan, M.; See, H. B.; Seeber, R. M.; Eidne, K. A.; Pfleger, K. D. Improvements to the Bioluminescence Resonance Energy Transfer (BRET) Technology for the Monitoring of G Protein-Coupled Receptors in Live Cells. J. Biomol. Screening 2008, 13, 888−898. (252) Bertrand, L.; Parent, S.; Caron, M.; Legault, M.; Joly, E.; Angers, S.; Bouvier, M.; Brown, M.; Houle, B.; Menard, L. The BRET2/Arrestin Assay in Stable Recombinant Cells: A Platform to Screen for Compounds that Interact with G Protein-Coupled Receptors (GPCRs). J. Recept. Signal Transduction Res. 2002, 22, 533−541. (253) Hamdan, F. F.; Audet, M.; Garneau, P.; Pelletier, J.; Bouvier, M. High-Throughput Screening of G Protein-Coupled Receptor Antagonists Using a Bioluminescence Resonance Energy Transfer 1Based β-Arrestin2 Recruitment Assay. J. Biomol. Screening 2005, 10, 463−475. (254) Kocan, M.; Dalrymple, M. B.; Seeber, R. M.; Feldman, B. J.; Pfleger, K. D. Enhanced BRET Technology for the Monitoring of Agonist-Induced and Agonist-Independent Interactions between GPCRs and β-Arrestins. Front. Endocrinol. 2010, 1, 12. (255) Charest, P. G.; Terrillon, S.; Bouvier, M. Monitoring AgonistPromoted Conformational Changes of β-Arrestin in Living Cells by Intramolecular BRET. EMBO Rep. 2005, 6, 334−340. (256) Shukla, A. K.; Violin, J. D.; Whalen, E. J.; Gesty-Palmer, D.; Shenoy, S. K.; Lefkowitz, R. J. Distinct Conformational Changes in βArrestin Report Biased Agonism at Seven-Transmembrane Receptors. Proc. Natl. Acad. Sci. U. S. A. 2008, 105, 9988−9993. (257) Lee, M. H.; Appleton, K. M.; Strungs, E. G.; Kwon, J. Y.; Morinelli, T. A.; Peterson, Y. K.; Laporte, S. A.; Luttrell, L. M. The Conformational Signature of Beta-arrestin2 Predicts Its Trafficking and Signalling Functions. Nature 2016, 531, 665−668. (258) Nuber, S.; Zabel, U.; Lorenz, K.; Nuber, A.; Milligan, G.; Tobin, A. B.; Lohse, M. J.; Hoffmann, C. Beta-Arrestin Biosensors Reveal a Rapid, Receptor-Dependent Activation/Deactivation Cycle. Nature 2016, 531, 661−664. (259) Gales, C.; Rebois, R. V.; Hogue, M.; Trieu, P.; Breit, A.; Hebert, T. E.; Bouvier, M. Real-Time Monitoring of Receptor and G Protein Interactions in Living Cells. Nat. Methods 2005, 2, 177−184. (260) Violin, J. D.; Lefkowitz, R. J. β-Arrestin-Biased Ligands at Seven-Transmembrane Receptors. Trends Pharmacol. Sci. 2007, 28, 416−422. (261) Masri, B.; Salahpour, A.; Didriksen, M.; Ghisi, V.; Beaulieu, J. M.; Gainetdinov, R. R.; Caron, M. G. Antagonism of Dopamine D2 Receptor/β-Arrestin 2 Interaction Is a Common Property of Clinically Effective Antipsychotics. Proc. Natl. Acad. Sci. U. S. A. 2008, 105, 13656−13661. (262) Griffin, B. A.; Adams, S. R.; Tsien, R. Y. Specific Covalent Labeling of Recombinant Protein Molecules inside Live Cells. Science 1998, 281, 269−272. (263) Adams, S. R.; Campbell, R. E.; Gross, L. A.; Martin, B. R.; Walkup, G. K.; Yao, Y.; Llopis, J.; Tsien, R. Y. New Biarsenical Ligands and Tetracysteine Motifs for Protein Labeling in Vitro and in Vivo: Synthesis and Biological Applications. J. Am. Chem. Soc. 2002, 124, 6063−6076. (264) Martin, B. R.; Giepmans, B. N.; Adams, S. R.; Tsien, R. Y. Mammalian Cell-Based Optimization of the Biarsenical-Binding Tetracysteine Motif for Improved Fluorescence and Affinity. Nat. Biotechnol. 2005, 23, 1308−1314. (265) Hoffmann, C.; Gaietta, G.; Zurn, A.; Adams, S. R.; Terrillon, S.; Ellisman, M. H.; Tsien, R. Y.; Lohse, M. J. Fluorescent Labeling of Tetracysteine-Tagged Proteins in Intact Cells. Nat. Protoc. 2010, 5, 1666−1677.

(266) Hoffmann, C.; Gaietta, G.; Bunemann, M.; Adams, S. R.; Oberdorff-Maass, S.; Behr, B.; Vilardaga, J. P.; Tsien, R. Y.; Ellisman, M. H.; Lohse, M. J. A FlAsH-Based FRET Approach to Determine G Protein-Coupled Receptor Activation in Living Cells. Nat. Methods 2005, 2, 171−176. (267) Vilardaga, J. P.; Nikolaev, V. O.; Lorenz, K.; Ferrandon, S.; Zhuang, Z.; Lohse, M. J. Conformational Cross-Talk between α(2a)Adrenergic and μ-Opioid Receptors Controls Cell Signaling. Nat. Chem. Biol. 2008, 4, 126−131. (268) Halo, T. L.; Appelbaum, J.; Hobert, E. M.; Balkin, D. M.; Schepartz, A. Selective Recognition of Protein Tetraserine Motifs with a Cell-Permeable, Pro-Fluorescent Bis-Boronic Acid. J. Am. Chem. Soc. 2009, 131, 438−439. (269) Kim, K. K.; Escobedo, J. O.; St Luce, N. N.; Rusin, O.; Wong, D.; Strongin, R. M. Postcolumn HPLC Detection of Mono- and Oligosaccharides with a Chemosensor. Org. Lett. 2003, 5, 5007−5010. (270) Nonaka, H.; Tsukiji, S.; Ojida, A.; Hamachi, I. Non-Enzymatic Covalent Protein Labeling Using a Reactive Tag. J. Am. Chem. Soc. 2007, 129, 15777−15779. (271) Nonaka, H.; Fujishima, S. H.; Uchinomiya, S. H.; Ojida, A.; Hamachi, I. Selective Covalent Labeling of Tag-Fused GPCR Proteins on Live Cell Surface with a Synthetic Probe for Their Functional Analysis. J. Am. Chem. Soc. 2010, 132, 9301−9309. (272) Reinhardt, U.; Lotze, J.; Zernia, S.; Morl, K.; Beck-Sickinger, A. G.; Seitz, O. Peptide-Templated Acyl Transfer: A Chemical Method for the Labeling of Membrane Proteins on Live Cells. Angew. Chem., Int. Ed. 2014, 53, 10237−10241. (273) Reinhardt, U.; Lotze, J.; Morl, K.; Beck-Sickinger, A. G.; Seitz, O. Rapid Covalent Fluorescence Labeling of Membrane Proteins on Live Cells via Coiled-Coil Templated Acyl Transfer. Bioconjugate Chem. 2015, 26, 2106−2117. (274) Rashidian, M.; Dozier, J. K.; Distefano, M. D. Enzymatic Labeling of Proteins: Techniques and Approaches. Bioconjugate Chem. 2013, 24, 1277−1294. (275) Pober, J. S.; Iwanij, V.; Reich, E.; Stryer, L. TransglutaminaseCatalyzed Insertion of a Fluorescent Probe into the Protease-Sensitive Region of Rhodopsin. Biochemistry 1978, 17, 2163−2168. (276) Juillerat, A.; Gronemeyer, T.; Keppler, A.; Gendreizig, S.; Pick, H.; Vogel, H.; Johnsson, K. Directed Evolution of O6-AlkylguanineDNA Alkyltransferase for Efficient Labeling of Fusion Proteins with Small Molecules in Vivo. Chem. Biol. 2003, 10, 313−317. (277) Keppler, A.; Gendreizig, S.; Gronemeyer, T.; Pick, H.; Vogel, H.; Johnsson, K. A General Method for the Covalent Labeling of Fusion Proteins with Small Molecules in Vivo. Nat. Biotechnol. 2002, 21, 86−89. (278) Keppler, A.; Pick, H.; Arrivoli, C.; Vogel, H.; Johnsson, K. Labeling of Fusion Proteins with Synthetic Fluorophores in Live Cells. Proc. Natl. Acad. Sci. U. S. A. 2004, 101, 9955−9959. (279) Gautier, A.; Juillerat, A.; Heinis, C.; Correa, I. R., Jr.; Kindermann, M.; Beaufils, F.; Johnsson, K. An Engineered Protein Tag for Multiprotein Labeling in Living Cells. Chem. Biol. 2008, 15, 128− 136. (280) Keppler, A.; Arrivoli, C.; Sironi, L.; Ellenberg, J. Fluorophores for Live Cell Imaging of AGT Fusion Proteins across the Visible Spectrum. BioTechniques 2006, 41, 167−170. (281) Lukinavicius, G.; Umezawa, K.; Olivier, N.; Honigmann, A.; Yang, G. Y.; Plass, T.; Mueller, V.; Reymond, L.; Correa, I. R.; Luo, Z. G.; et al. A Near-Infrared Fluorophore for Live-Cell Super-Resolution Microscopy of Cellular Proteins. Nat. Chem. 2013, 5, 132−139. (282) Maurel, D.; Comps-Agrar, L.; Brock, C.; Rives, M.-L.; Bourrier, E.; Ayoub, M. A.; Bazin, H.; Tinel, N.; Durroux, T.; Prezeau, L.; et al. Cell-Surface Protein-Protein Interaction Analysis with Time-Resolved Fret and SNAP-Tag Technologies: Application to GPCR Oligomerization. Nat. Methods 2008, 5, 561−567. (283) Petershans, A.; Wedlich, D.; Fruk, L. Bioconjugation of CdSe/ ZnS Nanoparticles with SNAP Tagged Proteins. Chem. Commun. 2011, 47, 10671−10673. AW

DOI: 10.1021/acs.chemrev.6b00084 Chem. Rev. XXXX, XXX, XXX−XXX

Chemical Reviews

Review

Chemotactic Receptor in the Plasma Membrane. Biochim. Biophys. Acta, Biomembr. 2011, 1808, 1701−1708. (303) Snaar-Jagalska, B. E.; Cambi, A.; Schmidt, T.; de Keijzer, S. Single-Molecule Imaging Technique to Study the Dynamic Regulation of GPCR Function at the Plasma Membrane. Methods Enzymol. 2013, 521, 47−67. (304) Suzuki, K. G. N.; Kasai, R. S.; Fujiwara, T. K.; Kusumi, A. Single-Molecule Imaging of Receptor-Receptor Interactions. Methods Cell Biol. 2013, 117, 373−390. (305) Latty, S. L.; Felce, J. H.; Weimann, L.; Lee, S. F.; Davis, S. J.; Klenerman, D. Referenced Single-Molecule Measurements Differentiate between GPCR Oligomerization States. Biophys. J. 2015, 109, 1798−1806. (306) Komatsuzaki, A.; Ohyanagi, T.; Tsukasaki, Y.; Miyanaga, Y.; Ueda, M.; Jin, T. Compact Halo-Ligand-Conjugated Quantum Dots for Multicolored Single-Molecule Imaging of Overcrowding GPCR Proteins on Cell Membranes. Small 2015, 11, 1396−1401. (307) Miller, L. W.; Cai, Y. F.; Sheetz, M. P.; Cornish, V. W. In Vivo Protein Labeling with Trimethoprim Conjugates: A Flexible Chemical Tag. Nat. Methods 2005, 2, 255−257. (308) Miller, L. W.; Cornish, V. W. Selective Chemical Labeling of Proteins in Living Cells. Curr. Opin. Chem. Biol. 2005, 9, 56−61. (309) Gallagher, S. S.; Sable, J. E.; Sheetz, M. P.; Cornish, V. W. An in Vivo Covalent TMP-Tag Based on Proximity-Induced Reactivity. ACS Chem. Biol. 2009, 4, 547−556. (310) Chen, Z.; Jing, C.; Gallagher, S. S.; Sheetz, M. P.; Cornish, V. W. Second-Generation Covalent TMP-Tag for Live Cell Imaging. J. Am. Chem. Soc. 2012, 134, 13692−13699. (311) Jing, C.; Cornish, V. W. A Fluorogenic TMP-Tag for High Signal-to-Background Intracellular Live Cell Imaging. ACS Chem. Biol. 2013, 8, 1704−1712. (312) Chen, I.; Howarth, M.; Lin, W. Y.; Ting, A. Y. Site-Specific Labeling of Cell Surface Proteins with Biophysical Probes Using Biotin Ligase. Nat. Methods 2005, 2, 99−104. (313) Fernandez-Suarez, M.; Baruah, H.; Martinez-Hernandez, L.; Xie, K. T.; Baskin, J. M.; Bertozzi, C. R.; Ting, A. Y. Redirecting Lipoic Acid Ligase for Cell Surface Protein Labeling with Small-Molecule Probes. Nat. Biotechnol. 2007, 25, 1483−1487. (314) Puthenveetil, S.; Liu, D. S.; White, K. A.; Thompson, S.; Ting, A. Y. Yeast Display Evolution of a Kinetically Efficient 13-Amino Acid Substrate for Lipoic Acid Ligase. J. Am. Chem. Soc. 2009, 131, 16430− 16438. (315) Chen, I.; Choi, Y. A.; Ting, A. Y. Phage Display Evolution of a Peptide Substrate for Yeast Biotin Ligase and Application to TwoColor Quantum Dot Labeling of Cell Surface Proteins. J. Am. Chem. Soc. 2007, 129, 6619−6625. (316) Slavoff, S. A.; Chen, I.; Choi, Y. A.; Ting, A. A. Y. Expanding the Substrate Tolerance of Biotin Ligase through Exploration of Enzymes from Diverse Species. J. Am. Chem. Soc. 2008, 130, 1160− 1162. (317) Roux, K. J.; Kim, D. I.; Raida, M.; Burke, B. A Promiscuous Biotin Ligase Fusion Protein Identifies Proximal and Interacting Proteins in Mammalian Cells. J. Cell Biol. 2012, 196, 801−810. (318) Fernandez-Suarez, M.; Chen, T. S.; Ting, A. Y. Protein-Protein Interaction Detection in Vitro and in Cells by Proximity Biotinylation. J. Am. Chem. Soc. 2008, 130, 9251−9253. (319) Cohen, J. D.; Zou, P.; Ting, A. Y. Site-Specific Protein Modification Using Lipoic Acid Ligase and Bis-Aryl Hydrazone Formation. ChemBioChem 2012, 13, 888−894. (320) Baruah, H.; Puthenveetil, S.; Choi, Y. A.; Shah, S.; Ting, A. Y. An Engineered Aryl Azide Ligase for Site-Specific Mapping of ProteinProtein Interactions through Photo-Cross-Linking. Angew. Chem., Int. Ed. 2008, 47, 7018−7021. (321) Uttamapinant, C.; White, K. A.; Baruah, H.; Thompson, S.; Fernandez-Suarez, M.; Puthenveetil, S.; Ting, A. Y. A Fluorophore Ligase for Site-Specific Protein Labeling inside Living Cells. Proc. Natl. Acad. Sci. U. S. A. 2010, 107, 10914−10919.

(284) Gronemeyer, T.; Godin, G.; Johnsson, K. Adding Value to Fusion Proteins through Covalent Labeling. Curr. Opin. Biotechnol. 2005, 16, 453−458. (285) Böhme, I.; Morl, K.; Bamming, D.; Meyer, C.; Beck-Sickinger, A. G. Tracking of Human Y Receptors in Living Cells -A Fluorescence Approach. Peptides 2007, 28, 226−234. (286) Roed, S. N.; Wismann, P.; Underwood, C. R.; Kulahin, N.; Iversen, H.; Cappelen, K. A.; Schaffer, L.; Lehtonen, J.; HecksherSoerensen, J.; Secher, A.; et al. Real-Time Trafficking and Signaling of the Glucagon-Like Peptide-1 Receptor. Mol. Cell. Endocrinol. 2014, 382, 938−949. (287) Ward, R. J.; Xu, T.-R.; Milligan, G. GPCR Oligomerization and Receptor Trafficking. Methods Enzymol. 2013, 521, 69−90. (288) Landgraf, D.; Okumus, B.; Chien, P.; Baker, T. A.; Paulsson, J. Segregation of Molecules at Cell Division Reveals Native Protein Localization. Nat. Methods 2012, 9, 480−482. (289) Zwier, J. M.; Bazin, H.; Lamarque, L.; Mathis, G. Luminescent Lanthanide Cryptates: From the Bench to the Bedside. Inorg. Chem. 2014, 53, 1854−1866. (290) Yuan, J. L.; Wang, G. L. Lanthanide Complex-Based Fluorescence Label for Time-Resolved Fluorescence Bioassay. J. Fluoresc. 2005, 15, 559−568. (291) Comps-Agrar, L.; Maurel, D.; Rondard, P.; Pin, J.-P.; Trinquet, E.; Prézeau, L. Cell-Surface Protein−Protein Interaction Analysis with Time-Resolved FRET and SNAP-Tag Technologies. Methods Mol. Biol. 2011, 756, 201−214. (292) Appelbe, S.; Milligan, G. Hetero-Oligomerization of Chemokine Receptors. Methods Enzymol. 2009, 461, 207−225. (293) Alvarez-Curto, E.; Ward, R. J.; Pediani, J. D.; Milligan, G. Ligand Regulation of the Quaternary Organization of Cell Surface M3Muscarinic Acetylcholine Receptors Analyzed by Fluorescence Resonance Energy Transfer (FRET) Imaging and Homogeneous Time-Resolved FRET. J. Biol. Chem. 2010, 285, 23318−23330. (294) Calebiro, D.; Rieken, F.; Wagner, J.; Sungkaworn, T.; Zabel, U.; Borzi, A.; Cocucci, E.; Zurn, A.; Lohse, M. J. Single-Molecule Analysis of Fluorescently Labeled G Protein-Coupled Receptors Reveals Complexes with Distinct Dynamics and Organization. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, 743−748. (295) Olofsson, L.; Felekyan, S.; Doumazane, E.; Scholler, P.; Fabre, L.; Zwier, J. M.; Rondard, P.; Seidel, C. A. M.; Pin, J.-P.; Margeat, E. Fine Tuning of Sub-Millisecond Conformational Dynamics Controls Metabotropic Glutamate Receptors Agonist Efficacy. Nat. Commun. 2014, 5, 5206. (296) Ward, R. J.; Pediani, J. D.; Milligan, G. Heteromultimerization of Cannabinoid CB(1) Receptor and Orexin Ox(1) Receptor Generates a Unique Complex in Which Both Protomers Are Regulated by Orexin A. J. Biol. Chem. 2011, 286, 37414−37428. (297) Doumazane, E.; Scholler, P.; Zwier, J. M.; Trinquet, E.; Rondard, P.; Pin, J. P. A New Approach to Analyze Cell Surface Protein Complexes Reveals Specific Heterodimeric Metabotropic Glutamate Receptors. FASEB J. 2011, 25, 66−77. (298) Kern, A.; Albarran-Zeckler, R.; Walsh, H. E.; Smith, R. G. ApoGhrelin Receptor Forms Heteromers with DRD2 in Hypothalamic Neurons and Is Essential for Anorexigenic Effects of DRD2 Agonism. Neuron 2012, 73, 317−332. (299) Pou, C.; la Cour, C. M.; Stoddart, L. A.; Millan, M. J.; Milligan, G. Functional Homomers and Heteromers of Dopamine D-2L and D3 Receptors Co-exist at the Cell Surface. J. Biol. Chem. 2012, 287, 8864−8878. (300) Los, G. V.; Encell, L. P.; McDougall, M. G.; Hartzell, D. D.; Karassina, N.; Zimprich, C.; Wood, M. G.; Learish, R.; Ohana, R. F.; Urh, M.; et al. HaloTag: A Novel Protein Labeling Technology for Cell Imaging and Protein Analysis. ACS Chem. Biol. 2008, 3, 373−382. (301) Locatelli-Hoops, S.; Sheen, F. C.; Zoubak, L.; Gawrisch, K.; Yeliseev, A. A. Application of Halo Tag Technology to Expression and Purification of Cannabinoid Receptor CB2. Protein Expression Purif. 2013, 89, 62−72. (302) de Keijzer, S.; Galloway, J.; Harms, G. S.; Devreotes, P. N.; Iglesias, P. A. Disrupting Microtubule Network Immobilizes Amoeboid AX

DOI: 10.1021/acs.chemrev.6b00084 Chem. Rev. XXXX, XXX, XXX−XXX

Chemical Reviews

Review

(322) Cohen, J. D.; Thompson, S.; Ting, A. Y. Structure-Guided Engineering of a Pacific Blue Fluorophore Ligase for Specific Protein Imaging in Living Cells. Biochemistry 2011, 50, 8221−8225. (323) Liu, D. S.; Nivon, L. G.; Richter, F.; Goldman, P. J.; Deerinck, T. J.; Yao, J. Z.; Richardson, D.; Phipps, W. S.; Ye, A. Z.; Ellisman, M. H.; et al. Computational Design of a Red Fluorophore Ligase for SiteSpecific Protein Labeling in Living Cells. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, E4551−4559. (324) Popp, M. W.; Antos, J. M.; Grotenbreg, G. M.; Spooner, E.; Ploegh, H. L. Sortagging: A Versatile Method for Protein Labeling. Nat. Chem. Biol. 2007, 3, 707−708. (325) Esteban, A.; Popp, M. W.; Vyas, V. K.; Strijbis, K.; Ploegh, H. L.; Fink, G. R. Fungal Recognition Is Mediated by the Association of Dectin-1 and Galectin-3 in Macrophages. Proc. Natl. Acad. Sci. U. S. A. 2011, 108, 14270−14275. (326) Hirota, N.; Yasuda, D.; Hashidate, T.; Yamamoto, T.; Yamaguchi, S.; Nagamune, T.; Nagase, T.; Shimizu, T.; Nakamura, M. Amino Acid Residues Critical for Endoplasmic Reticulum Export and Trafficking of Platelet-activating Factor Receptor. J. Biol. Chem. 2010, 285, 5931−5940. (327) Nakamura, M.; Yasuda, D.; Hirota, N.; Yamamoto, T.; Yamaguchi, S.; Shimizu, T.; Nagamune, T. Amino Acid Residues of G Protein-Coupled Receptors Critical for Endoplasmic Reticulum Export and Trafficking. Methods Enzymol. 2013, 521, 203−216. (328) Theile, C. S.; Witte, M. D.; Blom, A. E. M.; Kundrat, L.; Ploegh, H. L.; Guimaraes, C. P. Site-Specific N-Terminal Labeling of Proteins Using Sortase-Mediated Reactions. Nat. Protoc. 2013, 8, 1800−1807. (329) Guimaraes, C. P.; Witte, M. D.; Theile, C. S.; Bozkurt, G.; Kundrat, L.; Blom, A. E. M.; Ploegh, H. L. Site-Specific C-Terminal and Internal Loop Labeling of Proteins Using Sortase-Mediated Reactions. Nat. Protoc. 2013, 8, 1787−1799. (330) Schmidt, B.; Selmer, T.; Ingendoh, A.; Vonfigura, K. A Novel Amino-Acid Modification in Sulfatases That Is Defective in Multiple Sulfatase Deficiency. Cell 1995, 82, 271−278. (331) Carrico, I. S.; Carlson, B. L.; Bertozzi, C. R. Introducing Genetically Encoded Aldehydes into Proteins. Nat. Chem. Biol. 2007, 3, 321−322. (332) Rush, J. S.; Bertozzi, C. R. New Aldehyde Tag Sequences Identified by Screening Formylglycine Generating Enzymes in Vitro and in Vivo. J. Am. Chem. Soc. 2008, 130, 12240−12241. (333) Rabuka, D.; Rush, J. S.; deHart, G. W.; Wu, P.; Bertozzi, C. R. Site-Specific Chemical Protein Conjugation Using Genetically Encoded Aldehyde Tags. Nat. Protoc. 2012, 7, 1052−1067. (334) Rhee, H. W.; Zou, P.; Udeshi, N. D.; Martell, J. D.; Mootha, V. K.; Carr, S. A.; Ting, A. Y. Spatially Resolved Proteomic Mapping in Living Cells with the Engineered Peroxidase APEX2. Science 2013, 339, 1328−1331. (335) Lam, S. S.; Martell, J. D.; Kamer, K. J.; Deerinck, T. J.; Ellisman, M. H.; Mootha, V. K.; Ting, A. Y. Directed Evolution of APEX2 for Electron Microscopy and Proximity Labeling. Nat. Methods 2014, 12, 51−54. (336) Hung, V.; Udeshi, N. D.; Lam, S. S.; Loh, K. H.; Cox, K. J.; Pedram, K.; Carr, S. A.; Ting, A. Y. Spatially resolved proteomic mapping in living cells with the engineered peroxidase APEX2. Nat. Protoc. 2016, 11, 456−475. (337) Hung, V.; Zou, P.; Rhee, H. W.; Udeshi, N. D.; Cracan, V.; Svinkina, T.; Carr, S. A.; Mootha, V. K.; Ting, A. Y. Proteomic Mapping of the Human Mitochondrial Intermembrane Space in Live Cells via Ratiometric APEX Tagging. Mol. Cell 2014, 55, 332−341. (338) Martell, J. D.; Deerinck, T. J.; Sancak, Y.; Poulos, T. L.; Mootha, V. K.; Sosinsky, G. E.; Ellisman, M. H.; Ting, A. Y. Engineered Ascorbate Peroxidase as a Genetically Encoded Reporter for Electron Microscopy. Nat. Biotechnol. 2012, 30, 1143. (339) Howarth, M.; Ting, A. Y. Imaging Proteins in Live Mammalian Cells with Biotin Ligase and Monovalent Streptavidin. Nat. Protoc. 2008, 3, 534−545.

(340) Howarth, M.; Takao, K.; Hayashi, Y.; Ting, A. Y. Targeting Quantum Dots to Surface Proteins in Living Cells with Biotin Ligase. Proc. Natl. Acad. Sci. U. S. A. 2005, 102, 7583−7588. (341) Mize, G. J.; Harris, J. E.; Takayama, T. K.; Kulman, J. D. Regulated Expression of Active Biotinylated G Protein-Coupled Receptors in Mammalian Cells. Protein Expression Purif. 2008, 57, 280−289. (342) Schlinkmann, K. M.; Pluckthun, A. Directed Evolution of G Protein-Coupled Receptors for High Functional Expression and Detergent Stability. Methods Enzymol. 2013, 520, 67−97. (343) Slavoff, S. A.; Liu, D. S.; Cohen, J. D.; Ting, A. Y. Imaging Protein-Protein Interactions inside Living Cells Via InteractionDependent Fluorophore Ligation. J. Am. Chem. Soc. 2011, 133, 19769−19776. (344) Steel, E.; Murray, V. L.; Liu, A. P. Multiplex Detection of Homo- and Heterodimerization of G Protein-Coupled Receptors by Proximity Biotinylation. PLoS One 2014, 9, e93646. (345) Vilardaga, J. P.; Jean-Alphonse, F. G.; Gardella, T. J. Endosomal Generation of cAMP in GPCR Signaling. Nat. Chem. Biol. 2014, 10, 700−706. (346) Tsvetanova, N. G.; Irannejad, R.; von Zastrow, M. G Proteincoupled Receptor (GPCR) Signaling via Heterotrimeric G Proteins from Endosomes. J. Biol. Chem. 2015, 290, 6689−6696. (347) Benard, G.; Massa, F.; Puente, N.; Lourenco, J.; Bellocchio, L.; Soria-Gomez, E.; Matias, I.; Delamarre, A.; Metna-Laurent, M.; Cannich, A.; et al. Mitochondrial CB1 Receptors Regulate Neuronal Energy Metabolism. Nat. Neurosci. 2012, 15, 558−564. (348) Boutureira, O.; Bernardes, G. J. Advances in Chemical Protein Modification. Chem. Rev. 2015, 115, 2174−2195. (349) Sušac, L.; O’Connor, C.; Stevens, R. C.; Wüthrich, K. InMembrane Chemical Modification (IMCM) for Site-Specific Chromophore Labeling of GPCRs. Angew. Chem., Int. Ed. 2015, 54, 15246− 15249. (350) Chini, B.; Parenti, M. G protein-Coupled Receptors, Cholesterol and Palmitoylation: Facts about Fats. J. Mol. Endocrinol. 2009, 42, 371−379. (351) Wald, G.; Brown, P. K. The Role of Sulfhydryl Groups in the Bleaching and Synthesis of Rhodopsin. J. Gen. Physiol. 1952, 35, 797− 821. (352) Chen, Y. S.; Hubbell, W. L. Reactions of Sulfhydryl-Groups of Membrane-Bound Bovine Rhodopsin. Membr. Biochem. 1978, 1, 107− 130. (353) Karnik, S. S.; Khorana, H. G. Assembly of Functional Rhodopsin Requires a Disulfide Bond between Cysteine Residues 110 and 187. J. Biol. Chem. 1990, 265, 17520−17524. (354) Karnik, S.; Doi, T.; Molday, R.; Khorana, H. G. Expression of the Archaebacterial Bacterio-Opsin Gene with and without Signal Sequences in Escherichia Coli: The Expressed Proteins Are Located in the Membrane but Bind Retinal Poorly. Proc. Natl. Acad. Sci. U. S. A. 1990, 87, 8955−8959. (355) Ridge, K. D.; Lu, Z. J.; Liu, X.; Khorana, H. G. Structure and Function in Rhodopsin - Separation and Characterization of the Correctly Folded and Misfolded Opsins Produced on Expression of an Opsin Mutant-Gene Containing Only the Native Intradiscal Cysteine Codons. Biochemistry 1995, 34, 3261−3267. (356) Yang, K.; Farrens, D. L.; Hubbell, W. L.; Khorana, H. G. Structure and function in rhodopsin. Single cysteine substitution mutants in the cytoplasmic interhelical E-F loop region show positionspecific effects in transducin activation. Biochemistry 1996, 35, 12464− 12469. (357) Fraser, C. M. Site-Directed Mutagenesis of β-Adrenergic Receptors - Identification of Conserved Cysteine Residues That Independently Affect Ligand-Binding and Receptor Activation. J. Biol. Chem. 1989, 264, 9266−9270. (358) Kobilka, B.; Gether, U.; Seifert, R.; Lin, S.; Ghanouni, P. Examination of Ligand-Induced Conformational Changes in the β2Adrenergic Receptor. Life Sci. 1998, 62, 1509−1512. AY

DOI: 10.1021/acs.chemrev.6b00084 Chem. Rev. XXXX, XXX, XXX−XXX

Chemical Reviews

Review

(359) Rousselet, A.; Devaux, P. F. Interaction between Spin-Labeled Rhodopsin and Spin-Labeled Phospholipids in Retinal Outer Segment Disk Membranes. FEBS Lett. 1978, 93, 161−164. (360) Farahbakhsh, Z. T.; Altenbach, C.; Hubbell, W. L. Spin Labeled Cysteines as Sensors for Protein-Lipid Interaction and Conformation in Rhodopsin. Photochem. Photobiol. 1992, 56, 1019− 1033. (361) Resek, J. F.; Farahbakhsh, Z. T.; Hubbell, W. L.; Khorana, H. G. Formation of the Meta II Photointermediate Is Accompanied by Conformational Changes in the Cytoplasmic Surface of Rhodopsin. Biochemistry 1993, 32, 12025−12032. (362) Farahbakhsh, Z. T.; Ridge, K. D.; Khorana, H. G.; Hubbell, W. L. Mapping Light-Dependent Structural Changes in the Cytoplasmic Loop Connecting Helices C and D in Rhodopsin: a Site-Directed Spin Labeling Study. Biochemistry 1995, 34, 8812−8819. (363) Altenbach, C.; Yang, K.; Farrens, D. L.; Farahbakhsh, Z. T.; Khorana, H. G.; Hubbell, W. L. Structural Features and LightDependent Changes in the Cytoplasmic Interhelical E-F Loop Region of Rhodopsin: A Site-Directed Spin-Labeling Study. Biochemistry 1996, 35, 12470−12478. (364) Altenbach, C.; Klein-Seetharaman, J.; Hwa, J.; Khorana, H. G.; Hubbell, W. L. Structural Features and Light-Dependent Changes in the Sequence 59−75 Connecting Helices I and II in Rhodopsin: A Site-Directed Spin-Labeling Study. Biochemistry 1999, 38, 7945−7949. (365) Altenbach, C.; Cai, K.; Khorana, H. G.; Hubbell, W. L. Structural Features and Light-Dependent Changes in the Sequence 306−322 Extending from Helix VII to the Palmitoylation Sites in Rhodopsin: A Site-Directed Spin-Labeling Study. Biochemistry 1999, 38, 7931−7937. (366) Hubbell, W. L.; Altenbach, C.; Hubbell, C. M.; Khorana, H. G. Rhodopsin Structure, Dynamics, and Activation: A Perspective from Crystallography, Site-Directed Spin Labeling, Sulfhydryl Reactivity, and Disulfide Cross-Linking. Adv. Protein Chem. 2003, 63, 243−290. (367) Altenbach, C.; Kusnetzow, A. K.; Ernst, O. P.; Hofmann, K. P.; Hubbell, W. L. High-Resolution Distance Mapping in Rhodopsin Reveals the Pattern of Helix Movement Due to Activation. Proc. Natl. Acad. Sci. U. S. A. 2008, 105, 7439−7444. (368) Wu, C. W.; Stryer, L. Proximity Relationships in Rhodopsin. Proc. Natl. Acad. Sci. U. S. A. 1972, 69, 1104−1108. (369) Hargrave, P. A.; Mcdowell, J. H.; Curtis, D. R.; Wang, J. K.; Juszczak, E.; Fong, S. L.; Rao, J. K. M.; Argos, P. The Structure of Bovine Rhodopsin. Biophys. Struct. Mech. 1983, 9, 235−244. (370) Stryer, L. Fluorescence Energy-Transfer as a Spectroscopic Ruler. Annu. Rev. Biochem. 1978, 47, 819−846. (371) Farrens, D. L.; Khorana, H. G. Structure and Function in Rhodopsin: Measurement of the Rate of Metarhodopsin II Decay by Fluorescence Spectroscopy. J. Biol. Chem. 1995, 270, 5073−5076. (372) Schadel, S. A.; Heck, M.; Maretzki, D.; Filipek, S.; Teller, D. C.; Palczewski, K.; Hofmann, K. P. Ligand Channeling within a G ProteinCoupled Receptor: the Entry and Exit of Retinals in Native Opsin. J. Biol. Chem. 2003, 278, 24896−24903. (373) Gross, A. K.; Rao, V. R.; Oprian, D. D. Characterization of Rhodopsin Congenital Night Blindness Mutant T94I. Biochemistry 2003, 42, 2009−2015. (374) Piechnick, R.; Ritter, E.; Hildebrand, P. W.; Ernst, O. P.; Scheerer, P.; Hofmann, K. P.; Heck, M. Effect of Channel Mutations on the Uptake and Release of the Retinal Ligand in Opsin. Proc. Natl. Acad. Sci. U. S. A. 2012, 109, 5247−5252. (375) Sanchez-Martin, M. J.; Ramon, E.; Torrent-Burgues, J.; Garriga, P. Improved Conformational Stability of the Visual G Protein-Coupled Receptor Rhodopsin by Specific Interaction with Docosahexaenoic Acid Phospholipid. ChemBioChem 2013, 14, 639− 644. (376) Srinivasan, S.; Ramon, E.; Cordomi, A.; Garriga, P. Binding Specificity of Retinal Analogs to Photoactivated Visual Pigments Suggest Mechanism for Fine-Tuning GPCR-Ligand Interactions. Chem. Biol. 2014, 21, 369−378.

(377) Morrow, J. M.; Chang, B. S. Comparative Mutagenesis Studies of Retinal Release in Light-Activated Zebrafish Rhodopsin Using Fluorescence Spectroscopy. Biochemistry 2015, 54, 4507−4518. (378) Dunham, T. D.; Farrens, D. L. Conformational Changes in Rhodopsin: Movement of Helix F Detected by Site-Specific Chemical Labeling and Fluorescence Spectroscopy. J. Biol. Chem. 1999, 274, 1683−1690. (379) Imamoto, Y.; Kataoka, M.; Tokunaga, F.; Palczewski, K. LightInduced Conformational Changes of Rhodopsin Probed by Fluorescent Alexa594 Immobilized on the Cytoplasmic Surface. Biochemistry 2000, 39, 15225−15233. (380) Mielke, T.; Alexiev, U.; Glasel, M.; Otto, H.; Heyn, M. P. Light-Induced Changes in the Structure and Accessibility of the Cytoplasmic Loops of Rhodopsin in the Activated MII State. Biochemistry 2002, 41, 7875−7884. (381) Janz, J. M.; Farrens, D. L. Rhodopsin Activation Exposes a Key Hydrophobic Binding Site for the Transducin α-Subunit C Terminus. J. Biol. Chem. 2004, 279, 29767−29773. (382) Hoersch, D.; Otto, H.; Wallat, I.; Heyn, M. P. Monitoring the Conformational Changes of Photoactivated Rhodopsin from Microseconds to Seconds by Transient Fluorescence Spectroscopy. Biochemistry 2008, 47, 11518−11527. (383) Mansoor, S. E.; DeWitt, M. A.; Farrens, D. L. Distance Mapping in Proteins Using Fluorescence Spectroscopy: the Tryptophan-Induced Quenching (TriQ) Method. Biochemistry 2010, 49, 9722−9731. (384) Brunette, A. M. J.; Farrens, D. L. Distance Mapping in Proteins Using Fluorescence Spectroscopy: Tyrosine, Like Tryptophan, Quenches Bimane Fluorescence in a Distance-Dependent Manner. Biochemistry 2014, 53, 6290−6301. (385) Tsukamoto, H.; Farrens, D. L. A Constitutively Activating Mutation Alters the Dynamics of a Key Conformational Change in a Ligand-Free GPCR. J. Biol. Chem. 2013, 288, 28207−28216. (386) Yao, X.; Parnot, C.; Deupi, X.; Ratnala, V. R.; Swaminath, G.; Farrens, D.; Kobilka, B. Coupling Ligand Structure to Specific Conformational Switches in the β2-Adrenoceptor. Nat. Chem. Biol. 2006, 2, 417−422. (387) Peleg, G.; Ghanouni, P.; Kobilka, B. K.; Zare, R. N. SingleMolecule Spectroscopy of the β(2) Adrenergic Receptor: Observation of Conformational Substates in a Membrane Protein. Proc. Natl. Acad. Sci. U. S. A. 2001, 98, 8469−8474. (388) Lamichhane, R.; Liu, J. J.; Pljevaljcic, G.; White, K. L.; van der Schans, E.; Katritch, V.; Stevens, R. C.; Wuthrich, K.; Millar, D. P. Single-Molecule View of Basal Activity and Activation Mechanisms of the G Protein-Coupled Receptor β2AR. Proc. Natl. Acad. Sci. U. S. A. 2015, 112, 14254−14259. (389) Nygaard, R.; Zou, Y. Z.; Dror, R. O.; Mildorf, T. J.; Arlow, D. H.; Manglik, A.; Pan, A. C.; Liu, C. W.; Fung, J. J.; Bokoch, M. P.; et al. The Dynamic Process of β(2)-Adrenergic Receptor Activation. Cell 2013, 152, 532−542. (390) Klein-Seetharaman, J.; Getmanova, E. V.; Loewen, M. C.; Reeves, P. J.; Khorana, H. G. NMR Spectroscopy in Studies of LightInduced Structural Changes in Mammalian Rhodopsin: Applicability of Solution F-19 NMR. Proc. Natl. Acad. Sci. U. S. A. 1999, 96, 13744− 13749. (391) Loewen, M. C.; Klein-Seetharaman, J.; Getmanova, E. V.; Reeves, P. J.; Schwalbe, H.; Khorana, H. G. Solution 19F Nuclear Overhauser Effects in Structural Studies of the Cytoplasmic Domain of Mammalian Rhodopsin. Proc. Natl. Acad. Sci. U. S. A. 2001, 98, 4888− 4892. (392) Getmanova, E.; Patel, A. B.; Klein-Seetharaman, J.; Loewen, M. C.; Reeves, P. J.; Friedman, N.; Sheves, M.; Smith, S. O.; Khorana, H. G. NMR Spectroscopy of Phosphorylated Wild-Type Rhodopsin: Mobility of the Phosphorylated C-Terminus of Rhodopsin in the Dark and Upon Light Activation. Biochemistry 2004, 43, 1126−1133. (393) Liu, J. J.; Horst, R.; Katritch, V.; Stevens, R. C.; Wuthrich, K. Biased Signaling Pathways in β2-Adrenergic Receptor Characterized by 19F-NMR. Science 2012, 335, 1106−1110. AZ

DOI: 10.1021/acs.chemrev.6b00084 Chem. Rev. XXXX, XXX, XXX−XXX

Chemical Reviews

Review

(394) Manglik, A.; Kim, T. H.; Masureel, M.; Altenbach, C.; Yang, Z.; Hilger, D.; Lerch, M. T.; Kobilka, T. S.; Thian, F. S.; Hubbell, W. L.; et al. Structural Insights into the Dynamic Process of β2-Adrenergic Receptor Signaling. Cell 2015, 161, 1101−1111. (395) Ye, L.; Van Eps, N.; Zimmer, M.; Ernst, O. P.; Scott Prosser, R. Activation of the A adenosine G-protein-coupled receptor by conformational selection. Nature 2016, 533, 265. (396) Liapakis, G. Obtaining Structural and Functional Information for Gpcrs Using the Substituted-Cysteine Accessibility Method (SCAM). Curr. Pharm. Biotechnol. 2014, 15, 980−986. (397) Javitch, J. A.; Shi, L.; Liapakis, G. Use of the Substituted Cysteine Accessibility Method to Study the Structure and Function of G Protein-Coupled Receptors. Methods Enzymol. 2002, 343, 137−156. (398) Kahsai, A. W.; Xiao, K.; Rajagopal, S.; Ahn, S.; Shukla, A. K.; Sun, J.; Oas, T. G.; Lefkowitz, R. J. Multiple Ligand-Specific Conformations of the β(2)-Adrenergic Receptor. Nat. Chem. Biol. 2011, 7, 692−700. (399) Kahsai, A. W.; Rajagopal, S.; Sun, J. P.; Xiao, K. H. Monitoring Protein Conformational Changes and Dynamics Using Stable-Isotope Labeling and Mass Spectrometry. Nat. Protoc. 2014, 9, 1301−1319. (400) Bokoch, M. P.; Zou, Y.; Rasmussen, S. G.; Liu, C. W.; Nygaard, R.; Rosenbaum, D. M.; Fung, J. J.; Choi, H. J.; Thian, F. S.; Kobilka, T. S.; et al. Ligand-Specific Regulation of the Extracellular Surface of a GProtein-Coupled Receptor. Nature 2010, 463, 108−112. (401) Cordomi, A.; Gomez-Tamayo, J. C.; Gigoux, V.; Fourmy, D. Sulfur-Containing Amino Acids in 7TMRs: Molecular Gears for Pharmacology and Function. Trends Pharmacol. Sci. 2013, 34, 320− 331. (402) Resh, M. D. Palmitoylation of Ligands, Receptors, and Intracellular Signaling Molecules. Sci. Signal. 2006, 2006, re14. (403) Pless, S. A.; Ahern, C. A. Unnatural Amino Acids as Probes of Ligand-Receptor Interactions and Their Conformational Consequences. Annu. Rev. Pharmacol. Toxicol. 2013, 53, 211−229. (404) Noren, C. J.; Anthonycahill, S. J.; Griffith, M. C.; Schultz, P. G. A General-Method for Site-Specific Incorporation of Unnatural Amino-Acids into Proteins. Science 1989, 244, 182−188. (405) Goodman, H. M.; Abelson, J.; Landy, A.; Brenner, S.; Smith, J. D. Amber Suppression: A Nucleotide Change in the Anticodon of a Tyrosine Transfer RNA. Nature 1968, 217, 1019−1024. (406) Liebman, S. W.; Sherman, F.; Stewart, J. W. Isolation and Characterization of Amber Suppressors in Yeast. Genetics 1976, 82, 251−272. (407) Gesteland, R. F.; Wolfner, M.; Grisafi, P.; Fink, G.; Botstein, D.; Roth, J. R. Yeast Suppressors of UAA and UAG Nonsense Codons Work Efficiently in Vitro Via tRNA. Cell 1976, 7, 381−390. (408) Zhouravleva, G.; Frolova, L.; Le Goff, X.; Le Guellec, R.; IngeVechtomov, S.; Kisselev, L.; Philippe, M. Termination of Translation in Eukaryotes Is Governed by Two Interacting Polypeptide Chain Release Factors, eRF1 and eRF3. EMBO J. 1995, 14, 4065−4072. (409) Song, H.; Mugnier, P.; Das, A. K.; Webb, H. M.; Evans, D. R.; Tuite, M. F.; Hemmings, B. A.; Barford, D. The Crystal Structure of Human Eukaryotic Release Factor eRF1–Mechanism of Stop Codon Recognition and Peptidyl-tRNA Hydrolysis. Cell 2000, 100, 311−321. (410) Robertson, S. A.; Noren, C. J.; Anthony-Cahill, S. J.; Griffith, M. C.; Schultz, P. G. The Use of 5′-Phospho-2 Deoxyribocytidylylriboadenosine as a Facile Route to Chemical Aminoacylation of tRNA. Nucleic Acids Res. 1989, 17, 9649−9660. (411) Robertson, S. A.; Ellman, J. A.; Schultz, P. G. A General and Efficient Route for Chemical Aminoacylation of Transfer-RNAs. J. Am. Chem. Soc. 1991, 113, 2722−2729. (412) Mendel, D.; Ellman, J. A.; Schultz, P. G. Construction of a Light-Activated Protein by Unnatural Amino-Acid Mutagenesis. J. Am. Chem. Soc. 1991, 113, 2758−2760. (413) Judice, J. K.; Gamble, T. R.; Murphy, E. C.; Devos, A. M.; Schultz, P. G. Probing the Mechanism of Staphylococcal Nuclease with Unnatural Amino-Acids - Kinetic and Structural Studies. Science 1993, 261, 1578−1581.

(414) Kimata, Y.; Shimada, H.; Hirose, T.; Ishimura, Y. Role of Thr252 in Cytochrome P450CAM - a Study with Unnatural Amino-Acid Mutagenesis. Biochem. Biophys. Res. Commun. 1995, 208, 96−102. (415) Nowak, M. W.; Kearney, P. C.; Sampson, J. R.; Saks, M. E.; Labarca, C. G.; Silverman, S. K.; Zhong, W.; Thorson, J. S.; Abelson, J. N.; Davidson, N.; et al. Nicotinic Receptor-Binding Site Probed with Unnatural Amino-Acid-Incorporation in Intact Cells. Science 1995, 268, 439−442. (416) Monahan, S. L.; Lester, H. A.; Dougherty, D. A. Site-Specific Incorporation of Unnatural Amino Acids into Receptors Expressed in Mammalian Cells. Chem. Biol. 2003, 10, 573−580. (417) Torrice, M. M.; Bower, K. S.; Lester, H. A.; Dougherty, D. A. Probing the Role of the Cation-Pi Interaction in the Binding Sites of GPCRs Using Unnatural Amino Acids. Proc. Natl. Acad. Sci. U. S. A. 2009, 106, 11919−11924. (418) Martin, R. P.; Sibler, A. P.; Dirheimer, G.; de Henau, S.; Grosjean, H. Yeast Mitochondrial tRNATrp Injected with E. Coli Activating Enzyme into Xenopus Oocytes Suppresses UGA Termination. Nature 1981, 293, 235−237. (419) Nakamura, Y.; Gojobori, T.; Ikemura, T. Codon Usage Tabulated from International DNA Sequence Databases: Status for the Year 2000. Nucleic Acids Res. 2000, 28, 292−292. (420) Liu, D. R.; Schultz, P. G. Progress toward the Evolution of an Organism with an Expanded Genetic Code. Proc. Natl. Acad. Sci. U. S. A. 1999, 96, 4780−4785. (421) Wang, L.; Magliery, T. J.; Liu, D. R.; Schultz, P. G. A New Functional Suppressor tRNA/Aminoacyl-tRNA Synthetase Pair for the in Vivo Incorporation of Unnatural Amino Acids into Proteins. J. Am. Chem. Soc. 2000, 122, 5010−5011. (422) Wang, L.; Schultz, P. G. A General Approach for the Generation of Orthogonal tRNAs. Chem. Biol. 2001, 8, 883−890. (423) Wang, L.; Brock, A.; Herberich, B.; Schultz, P. G. Expanding the Genetic Code of Escherichia Coli. Science 2001, 292, 498−500. (424) Sakamoto, K.; Hayashi, A.; Sakamoto, A.; Kiga, D.; Nakayama, H.; Soma, A.; Kobayashi, T.; Kitabatake, M.; Takio, K.; Saito, K.; et al. Site-Specific Incorporation of an Unnatural Amino Acid into Proteins in Mammalian Cells. Nucleic Acids Res. 2002, 30, 4692−4699. (425) Deiters, A.; Cropp, T. A.; Mukherji, M.; Chin, J. W.; Anderson, J. C.; Schultz, P. G. Adding Amino Acids with Novel Reactivity to the Genetic Code of Saccharomyces Cerevisiae. J. Am. Chem. Soc. 2003, 125, 11782−11783. (426) Chin, J. W.; Cropp, T. A.; Chu, S.; Meggers, E.; Schultz, P. G. Progress toward an Expanded Eukaryotic Genetic Code. Chem. Biol. 2003, 10, 511−519. (427) Chin, J. W.; Cropp, T. A.; Anderson, J. C.; Mukherji, M.; Zhang, Z. W.; Schultz, P. G. An Expanded Eukaryotic Genetic Code. Science 2003, 301, 964−967. (428) Shafer, A. M.; Kalai, T.; Bin Liu, S. Q.; Hideg, K.; Voss, J. C. Site-Specific Insertion of Spin-Labeled L-Amino Acids in Xenopus Oocytes. Biochemistry 2004, 43, 8470−8482. (429) Ye, S.; Riou, M.; Carvalho, S.; Paoletti, P. Expanding the Genetic Code in Xenopus Laevis Oocytes. ChemBioChem 2013, 14, 230−235. (430) Mukai, T.; Wakiyama, M.; Sakamoto, K.; Yokoyama, S. Genetic Encoding of Non-Natural Amino Acids in Drosophila Melanogaster Schneider 2 Cells. Protein Sci. 2010, 19, 440−448. (431) Liu, W.; Brock, A.; Chen, S.; Schultz, P. G. Genetic Incorporation of Unnatural Amino Acids into Proteins in Mammalian Cells. Nat. Methods 2007, 4, 239−244. (432) Ye, S.; Köhrer, C.; Huber, T.; Kazmi, M.; Sachdev, P.; Yan, E. C. Y.; Bhagat, A.; RajBhandary, U. L.; Sakmar, T. P. Site-Specific Incorporation of Keto Amino Acids into Functional G ProteinCoupled Receptors Using Unnatural Amino Acid Mutagenesis. J. Biol. Chem. 2008, 283, 1525−1533. (433) Wang, W.; Takimoto, J. K.; Louie, G. V.; Baiga, T. J.; Noel, J. P.; Lee, K. F.; Slesinger, P. A.; Wang, L. Genetically Encoding Unnatural Amino Acids for Cellular and Neuronal Studies. Nat. Neurosci. 2007, 10, 1063−1072. BA

DOI: 10.1021/acs.chemrev.6b00084 Chem. Rev. XXXX, XXX, XXX−XXX

Chemical Reviews

Review

(434) Shen, B.; Xiang, Z.; Miller, B.; Louie, G.; Wang, W.; Noel, J. P.; Gage, F. H.; Wang, L. Genetically Encoding Unnatural Amino Acids in Neural Stem Cells and Optically Reporting Voltage-Sensitive Domain Changes in Differentiated Neurons. Stem Cells 2011, 29, 1231−1240. (435) Anderson, J. C.; Schultz, P. G. Adaptation of an Orthogonal Archaeal Leucyl-tRNA and Synthetase Pair for Four-Base, Amber, and Opal Suppression. Biochemistry 2003, 42, 9598−9608. (436) Kowal, A. K.; Köhrer, C.; RajBhandary, U. L. Twenty-First Aminoacyl-tRNA Synthetase-Suppressor tRNA Pairs for Possible Use in Site-Specific Incorporation of Amino Acid Analogues into Proteins in Eukaryotes and in Eubacteria. Proc. Natl. Acad. Sci. U. S. A. 2001, 98, 2268−2273. (437) Santoro, S. W.; Anderson, J. C.; Lakshman, V.; Schultz, P. G. An Archaebacteria-Derived Glutamyl-tRNA Synthetase and tRNA Pair for Unnatural Amino Acid Mutagenesis of Proteins in Escherichia Coli. Nucleic Acids Res. 2003, 31, 6700−6709. (438) Chatterjee, A.; Xiao, H.; Yang, P. Y.; Soundararajan, G.; Schultz, P. G. A Tryptophanyl-tRNA Synthetase/tRNA Pair for Unnatural Amino Acid Mutagenesis in E. Coli. Angew. Chem., Int. Ed. 2013, 52, 5106−5109. (439) Polycarpo, C.; Ambrogelly, A.; Berube, A.; Winbush, S. A. M.; McCloskey, J. A.; Crain, P. F.; Wood, J. L.; Soll, D. An AminoacyltRNA Synthetase That Specifically Activates Pyrrolysine. Proc. Natl. Acad. Sci. U. S. A. 2004, 101, 12450−12454. (440) Blight, S. K.; Larue, R. C.; Mahapatra, A.; Longstaff, D. G.; Chang, E.; Zhao, G.; Kang, P. T.; Green-Church, K. B.; Chan, M. K.; Krzycki, J. A. Direct Charging of tRNA(CUA) with Pyrrolysine in Vitro and in Vivo. Nature 2004, 431, 333−335. (441) Mukai, T.; Kobayashi, T.; Hino, N.; Yanagisawa, T.; Sakamoto, K.; Yokoyama, S. Adding L-Lysine Derivatives to the Genetic Code of Mammalian Cells with Engineered Pyrrolysyl-tRNA Synthetases. Biochem. Biophys. Res. Commun. 2008, 371, 818−822. (442) Chen, P. R.; Groff, D.; Guo, J. T.; Ou, W. J.; Cellitti, S.; Geierstanger, B. H.; Schultz, P. G. A Facile System for Encoding Unnatural Amino Acids in Mammalian Cells. Angew. Chem., Int. Ed. 2009, 48, 4052−4055. (443) Chin, J. W. Expanding and Reprogramming the Genetic Code of Cells and Animals. Annu. Rev. Biochem. 2014, 83, 379−408. (444) Srinivasan, G.; James, C. M.; Krzycki, J. A. Pyrrolysine Encoded by Uag in Archaea: Charging of a UAG-Decoding Specialized tRNA. Science 2002, 296, 1459−1462. (445) Krzycki, J. A. The Direct Genetic Encoding of Pyrrolysine. Curr. Opin. Microbiol. 2005, 8, 706−712. (446) Guo, L. T.; Wang, Y. S.; Nakamura, A.; Eiler, D.; Kavran, J. M.; Wong, M.; Kiessling, L. L.; Steitz, T. A.; O’Donoghue, P.; Soll, D. Polyspecific Pyrrolysyl-tRNA Synthetases from Directed Evolution. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, 16724−16729. (447) Nozawa, K.; O’Donoghue, P.; Gundllapalli, S.; Araiso, Y.; Ishitani, R.; Umehara, T.; Soll, D.; Nureki, O. Pyrrolysyl-tRNA Synthetase-tRNA(Pyl) Structure Reveals the Molecular Basis of Orthogonality. Nature 2009, 457, 1163−1167. (448) Wan, W.; Tharp, J. M.; Liu, W. R. Pyrrolysyl-tRNA Synthetase: An Ordinary Enzyme but an Outstanding Genetic Code Expansion Tool. Biochim. Biophys. Acta, Proteins Proteomics 2014, 1844, 1059− 1070. (449) Greiss, S.; Chin, J. W. Expanding the Genetic Code of an Animal. J. Am. Chem. Soc. 2011, 133, 14196−14199. (450) Bianco, A.; Townsley, F. M.; Greiss, S.; Lang, K.; Chin, J. W. Expanding the Genetic Code of Drosophila Melanogaster. Nat. Chem. Biol. 2012, 8, 748−750. (451) Li, F.; Zhang, H.; Sun, Y.; Pan, Y.; Zhou, J.; Wang, J. Expanding the Genetic Code for Photoclick Chemistry in E. Coli, Mammalian Cells, and A. Thaliana. Angew. Chem., Int. Ed. 2013, 52, 9700−9704. (452) Elsässer, S. J.; Ernst, R. J.; Walker, O. S.; Chin, J. W. Genetic Code Expansion in Stable Cell Lines Enables Encoded Chromatin Modification. Nat. Methods 2016, 13, 158−164.

(453) Hendrickson, T. L.; de Crecy-Lagard, V.; Schimmel, P. Incorporation of Nonnatural Amino Acids into Proteins. Annu. Rev. Biochem. 2004, 73, 147−176. (454) Wang, L.; Xie, J.; Schultz, P. G. Expanding the Genetic Code. Annu. Rev. Biophys. Biomol. Struct. 2006, 35, 225−249. (455) Xie, J.; Schultz, P. G. A Chemical Toolkit for Proteins - An Expanded Genetic Code. Nat. Rev. Mol. Cell Biol. 2006, 7, 775−782. (456) Liu, C. C.; Schultz, P. G. Adding New Chemistries to the Genetic Code. Annu. Rev. Biochem. 2010, 79, 413−444. (457) Lang, K.; Chin, J. W. Cellular Incorporation of Unnatural Amino Acids and Bioorthogonal Labeling of Proteins. Chem. Rev. 2014, 114, 4764−4806. (458) Köhrer, C.; Yoo, J. H.; Bennett, M.; Schaack, J.; RajBhandary, U. L. A Possible Approach to Two Different Unnatural Site-Specific Insertion of Amino Acids into Proteins in Mammalian Cells via Nonsense Suppression. Chem. Biol. 2003, 10, 1095−1102. (459) Schmied, W. H.; Elsasser, S. J.; Uttamapinant, C.; Chin, J. W. Efficient Multisite Unnatural Amino Acid Incorporation in Mammalian Cells via Optimized Pyrrolysyl tRNA Synthetase/tRNA Expression and Engineered eRF1. J. Am. Chem. Soc. 2014, 136, 15577−15583. (460) Neumann, H.; Wang, K.; Davis, L.; Garcia-Alai, M.; Chin, J. W. Encoding Multiple Unnatural Amino Acids Via Evolution of a Quadruplet-Decoding Ribosome. Nature 2010, 464, 441−444. (461) Wan, W.; Huang, Y.; Wang, Z. Y.; Russell, W. K.; Pai, P. J.; Russell, D. H.; Liu, W. R. A Facile System for Genetic Incorporation of Two Different Noncanonical Amino Acids into One Protein in Escherichia Coli. Angew. Chem., Int. Ed. 2010, 49, 3211−3214. (462) Chatterjee, A.; Sun, S. B.; Furman, J. L.; Xiao, H.; Schultz, P. G. Versatile Platform for Single- and Multiple-Unnatural Amino Acid Mutagenesis in Escherichia Coli. Biochemistry 2013, 52, 1828−1837. (463) Wang, K.; Sachdeva, A.; Cox, D. J.; Wilf, N. W.; Lang, K.; Wallace, S.; Mehl, R. A.; Chin, J. W. Optimized Orthogonal Translation of Unnatural Amino Acids Enables Spontaneous Protein Double-Labelling and FRET. Nat. Chem. 2014, 6, 393−403. (464) Nikic, I.; Lemke, E. A. Genetic Code Expansion Enabled SiteSpecific Dual-Color Protein Labeling: Superresolution Microscopy and Beyond. Curr. Opin. Chem. Biol. 2015, 28, 164−173. (465) Damian, M.; Marie, J.; Leyris, J. P.; Fehrentz, J. A.; Verdie, P.; Martinez, J.; Baneres, J. L.; Mary, S. High Constitutive Activity Is an Intrinsic Feature of Ghrelin Receptor Protein a Study with a Functional Monomeric GHS-R1a Receptor Reconstituted in Lipid Discs. J. Biol. Chem. 2012, 287, 3630−3641. (466) Park, S. H.; Das, B. B.; Casagrande, F.; Tian, Y.; Nothnagel, H. J.; Chu, M. N.; Kiefer, H.; Maier, K.; De Angelis, A. A.; Marassi, F. M.; et al. Structure of the Chemokine Receptor CXCR1 in Phospholipid Bilayers. Nature 2012, 491, 779−783. (467) Wiktor, M.; Morin, S.; Sass, H. J.; Kebbel, F.; Grzesiek, S. Biophysical and Structural Investigation of Bacterially Expressed and Engineered CCR5, a G Protein-Coupled Receptor. J. Biomol. NMR 2013, 55, 79−95. (468) Huang, L. Y.; Umanah, G.; Hauser, M.; Son, C.; Arshava, B.; Naider, F.; Becker, J. M. Unnatural Amino Acid Replacement in a Yeast G Protein-Coupled Receptor in Its Native Environment. Biochemistry 2008, 47, 5638−5648. (469) Grunbeck, A.; Huber, T.; Abrol, R.; Trzaskowski, B.; Goddard, W. A.; Sakmar, T. P. Genetically Encoded Photo-Cross-Linkers Map the Binding Site of an Allosteric Drug on a G Protein-Coupled Receptor. ACS Chem. Biol. 2012, 7, 967−972. (470) Naganathan, S.; Ye, S.; Sakmar, T. P.; Huber, T. Site-Specific Epitope Tagging of G Protein-Coupled Receptors by Bioorthogonal Modification of a Genetically Encoded Unnatural Amino Acid. Biochemistry 2013, 52, 1028−1036. (471) Naganathan, S.; Ray-Saha, S.; Park, M.; Tian, H.; Sakmar, T. P.; Huber, T. Multiplex Detection of Functional G Protein-Coupled Receptors Harboring Site-Specifically Modified Unnatural Amino Acids. Biochemistry 2015, 54, 776−786. (472) Coin, I.; Katritch, V.; Sun, T. T.; Xiang, Z.; Siu, F. Y.; Beyermann, M.; Stevens, R. C.; Wang, L. Genetically Encoded BB

DOI: 10.1021/acs.chemrev.6b00084 Chem. Rev. XXXX, XXX, XXX−XXX

Chemical Reviews

Review

Chemical Probes in Cells Reveal the Binding Path of Urocortin-I to CRF Class B GPCR. Cell 2013, 155, 1258−1269. (473) Valentin-Hansen, L.; Park, M.; Huber, T.; Grunbeck, A.; Naganathan, S.; Schwartz, T. W.; Sakmar, T. P. Mapping Substance P Binding Sites on the Neurokinin-1 Receptor Using Genetic Incorporation of a Photoreactive Amino Acid. J. Biol. Chem. 2014, 289, 18045−18054. (474) Damian, M.; Mary, S.; Maingot, M.; M’Kadmi, C.; Gagne, D.; Leyris, J. P.; Denoyelle, S.; Gaibelet, G.; Gavara, L.; Costa, M. G. D.; et al. Ghrelin Receptor Conformational Dynamics Regulate the Transition from a Preassembled to an Active Receptor:Gq complex. Proc. Natl. Acad. Sci. U. S. A. 2015, 112, 1601−1606. (475) Cornish, V. W.; Benson, D. R.; Altenbach, C. A.; Hideg, K.; Hubbell, W. L.; Schultz, P. G. Site-Specific Incorporation of Biophysical Probes into Proteins. Proc. Natl. Acad. Sci. U. S. A. 1994, 91, 2910−2914. (476) Ye, S.; Huber, T.; Vogel, R.; Sakmar, T. P. FTIR Analysis of GPCR Activation Using Azido Probes. Nat. Chem. Biol. 2009, 5, 397− 399. (477) Ye, S.; Zaitseva, E.; Caltabiano, G.; Schertler, G. F.; Sakmar, T. P.; Deupi, X.; Vogel, R. Tracking G Protein-Coupled Receptor Activation Using Genetically Encoded Infrared Probes. Nature 2010, 464, 1386−1389. (478) Grunbeck, A.; Huber, T.; Sachdev, P.; Sakmar, T. P. Mapping the Ligand-Binding Site on a G Protein-Coupled Receptor (GPCR) Using Genetically Encoded Photocrosslinkers. Biochemistry 2011, 50, 3411−3413. (479) Naganathan, S.; Grunbeck, A.; Tian, H.; Huber, T.; Sakmar, T. P. Genetically-Encoded Molecular Probes to Study G Protein-Coupled Receptors. J. Vis. Exp. 2013, DOI: 10.3791/50588. (480) Grunbeck, A.; Sakmar, T. P. Probing G Protein-Coupled Receptor-Ligand Interactions with Targeted Photoactivatable CrossLinkers. Biochemistry 2013, 52, 8625−8632. (481) Coin, I.; Perrin, M. H.; Vale, W. W.; Wang, L. Photo-CrossLinkers Incorporated into G Protein-Coupled Receptors in Mammalian Cells: A Ligand Comparison. Angew. Chem., Int. Ed. 2011, 50, 8077−8081. (482) Ray-Saha, S.; Huber, T.; Sakmar, T. P. Antibody Epitopes on G Protein-Coupled Receptors Mapped with Genetically Encoded Photoactivatable Cross-Linkers. Biochemistry 2014, 53, 1302−1310. (483) Sato, S.; Mimasu, S.; Sato, A.; Hino, N.; Sakamoto, K.; Umehara, T.; Yokoyama, S. Crystallographic Study of a SiteSpecifically Cross-Linked Protein Complex with a Genetically Incorporated Photoreactive Amino Acid. Biochemistry 2011, 50, 250−257. (484) Xiang, Z.; Ren, H. Y.; Hu, Y. S.; Coin, I.; Wei, J.; Cang, H.; Wang, L. Adding an Unnatural Covalent Bond to Proteins through Proximity-Enhanced Bioreactivity. Nat. Methods 2013, 10, 885−888. (485) Tippmann, E. M.; Liu, W.; Summerer, D.; Mack, A. V.; Schultz, P. G. A Genetically Encoded Diazirine Photocrosslinker in Escherichia Coli. ChemBioChem 2007, 8, 2210−2214. (486) Chou, C. J.; Uprety, R.; Davis, L.; Chin, J. W.; Deiters, A. Genetically Encoding an Aliphatic Diazirine for Protein Photocrosslinking. Chem. Sci. 2011, 2, 480−483. (487) Lin, S. X.; Zhang, Z. R.; Xu, H.; Li, L.; Chen, S.; Li, J.; Hao, Z. Y.; Chen, P. R. Site-Specific Incorporation of Photo-Cross-Linker and Bioorthogonal Amino Acids into Enteric Bacterial Pathogens. J. Am. Chem. Soc. 2011, 133, 20581−20587. (488) Schultz, K. C.; Supekova, L.; Ryu, Y.; Xie, J.; Perera, R.; Schultz, P. G. A Genetically Encoded Infrared Probe. J. Am. Chem. Soc. 2006, 128, 13984−13985. (489) Gai, X. S.; Coutifaris, B. A.; Brewer, S. H.; Fenlon, E. E. A Direct Comparison of Azide and Nitrile Vibrational Probes. Phys. Chem. Chem. Phys. 2011, 13, 5926−5930. (490) Bazewicz, C. G.; Liskov, M. T.; Hines, K. J.; Brewer, S. H. Sensitive, Site-Specific, and Stable Vibrational Probe of Local Protein Environments: 4-Azidomethyl-L-Phenylalanine. J. Phys. Chem. B 2013, 117, 8987−8993.

(491) Jackson, J. C.; Hammill, J. T.; Mehl, R. A. Site-Specific Incorporation of a (19)F-Amino Acid into Proteins as an NMR Probe for Characterizing Protein Structure and Reactivity. J. Am. Chem. Soc. 2007, 129, 1160−1166. (492) Hammill, J. T.; Miyake-Stoner, S.; Hazen, J. L.; Jackson, J. C.; Mehl, R. A. Preparation of Site-Specifically Labeled Fluorinated Proteins for 19F-NMR Structural Characterization. Nat. Protoc. 2007, 2, 2601−2607. (493) Schmidt, M. J.; Borbas, J.; Drescher, M.; Summerer, D. A Genetically Encoded Spin Label for Electron Paramagnetic Resonance Distance Measurements. J. Am. Chem. Soc. 2014, 136, 1238−1241. (494) Tsao, M. L.; Summerer, D.; Ryu, Y. H.; Schultz, P. G. The Genetic Incorporation of a Distance Probe into Proteins in Escherichia Coli. J. Am. Chem. Soc. 2006, 128, 4572−4573. (495) Lv, X. X.; Yu, Y.; Zhou, M.; Hu, C.; Gao, F.; Li, J. S.; Liu, X. H.; Deng, K.; Zheng, P.; Gong, W. M.; et al. Ultrafast Photoinduced Electron Transfer in Green Fluorescent Protein Bearing a Genetically Encoded Electron Acceptor. J. Am. Chem. Soc. 2015, 137, 7270−7273. (496) Bose, M.; Groff, D.; Xie, J. M.; Brustad, E.; Schultz, P. G. The Incorporation of a Photoisomerizable Amino Acid into Proteins in E. Coli. J. Am. Chem. Soc. 2006, 128, 388−389. (497) Beharry, A. A.; Woolley, G. A. Azobenzene Photoswitches for Biomolecules. Chem. Soc. Rev. 2011, 40, 4422−4437. (498) Hoppmann, C.; Lacey, V. K.; Louie, G. V.; Wei, J.; Noel, J. P.; Wang, L. Genetically Encoding Photoswitchable Click Amino Acids in Escherichia Coli and Mammalian Cells. Angew. Chem., Int. Ed. 2014, 53, 3932−3936. (499) Turcatti, G.; Nemeth, K.; Edgerton, M. D.; Meseth, U.; Talabot, F.; Peitsch, M.; Knowles, J.; Vogel, H.; Chollet, A. Probing the Structure and Function of the Tachykinin Neurokinin-2 Receptor through Biosynthetic Incorporation of Fluorescent Amino Acids at Specific Sites. J. Biol. Chem. 1996, 271, 19991−19998. (500) Cohen, B. E.; McAnaney, T. B.; Park, E. S.; Jan, Y. N.; Boxer, S. G.; Jan, L. Y. Probing Protein Electrostatics with a Synthetic Fluorescent Amino Acid. Science 2002, 296, 1700−1703. (501) Ninomiya, K.; Kurita, T.; Hohsaka, T.; Sisido, M. Facile Aminoacylation of pdCpA Dinucleotide with a Nonnatural Amino Acid in Cationic Micelle. Chem. Commun. 2003, 2242−2243. (502) Kajihara, D.; Abe, R.; Iijima, I.; Komiyama, C.; Sisido, M.; Hohsaka, T. FRET Analysis of Protein Conformational Change through Position-Specific Incorporation of Fluorescent Amino Acids. Nat. Methods 2006, 3, 923−929. (503) Pantoja, R.; Rodriguez, E. A.; Dibas, M. I.; Dougherty, D. A.; Lester, H. A. Single-Molecule Imaging of a Fluorescent Unnatural Amino Acid Incorporated into Nicotinic Receptors. Biophys. J. 2009, 96, 226−237. (504) Zhang, Z. W.; Alfonta, L.; Tian, F.; Bursulaya, B.; Uryu, S.; King, D. S.; Schultz, P. G. Selective Incorporation of 5-Hydroxytryptophan into Proteins in Mammalian Cells. Proc. Natl. Acad. Sci. U. S. A. 2004, 101, 8882−8887. (505) Wang, J. Y.; Xie, J. M.; Schultz, P. G. A Genetically Encoded Fluorescent Amino Acid. J. Am. Chem. Soc. 2006, 128, 8738−8739. (506) Summerer, D.; Chen, S.; Wu, N.; Deiters, A.; Chin, J. W.; Schultz, P. G. A Genetically Encoded Fluorescent Amino Acid. Proc. Natl. Acad. Sci. U. S. A. 2006, 103, 9785−9789. (507) Lee, H. S.; Guo, J. T.; Lemke, E. A.; Dimla, R. D.; Schultz, P. G. Genetic Incorporation of a Small, Environmentally Sensitive, Fluorescent Probe into Proteins in Saccharomyces Cerevisiae. J. Am. Chem. Soc. 2009, 131, 12921−12923. (508) Chatterjee, A.; Guo, J.; Lee, H. S.; Schultz, P. G. A Genetically Encoded Fluorescent Probe in Mammalian Cells. J. Am. Chem. Soc. 2013, 135, 12540−12543. (509) Tian, H.; Sakmar, T. P.; Huber, T. Site-Specific Labeling of Genetically Encoded Azido Groups for Multicolor, Single-Molecule Fluorescence Imaging of GPCRs. Methods Cell Biol. 2013, 117, 267− 303. (510) Tian, H.; Naganathan, S.; Kazmi, M. A.; Schwartz, T. W.; Sakmar, T. P.; Huber, T. Bioorthogonal Fluorescent Labeling of BC

DOI: 10.1021/acs.chemrev.6b00084 Chem. Rev. XXXX, XXX, XXX−XXX

Chemical Reviews

Review

Functional G Protein-Coupled Receptors. ChemBioChem 2014, 15, 1820−1829. (511) Tian, H.; Sakmar, T. P.; Huber, T. Micelle-Enhanced Bioorthogonal Labeling of Genetically Encoded Azido Groups on the Lipid-Embedded Surface of a GPCR. ChemBioChem 2015, 16, 1314−1322. (512) Park, M.; Sivertsen, B. B.; Els-Heindl, S.; Huber, T.; Holst, B.; Beck-Sickinger, A. G.; Schwartz, T. W.; Sakmar, T. P. Bioorthogonal Labeling of Ghrelin Receptor to Facilitate Studies of LigandDependent Conformational Dynamics. Chem. Biol. 2015, 22, 1431− 1436. (513) Lang, K.; Chin, J. W. Bioorthogonal Reactions for Labeling Proteins. ACS Chem. Biol. 2014, 9, 16−20. (514) Cornish, V. W.; Hahn, K. M.; Schultz, P. G. Site-Specific Protein Modification Using a Ketone Handle. J. Am. Chem. Soc. 1996, 118, 8150−8151. (515) Hubbard, R. The Thermal Stability of Rhodopsin and Opsin. J. Gen. Physiol. 1958, 42, 259−280. (516) Huber, T.; Naganathan, S.; Tian, H.; Ye, S. X.; Sakmar, T. P. Unnatural Amino Acid Mutagenesis of GPCRs Using Amber Codon Suppression and Bioorthogonal Labeling. Methods Enzymol. 2013, 520, 281−305. (517) Grimsrud, P. A.; Xie, H.; Griffin, T. J.; Bernlohr, D. A. Oxidative Stress and Covalent Modification of Protein with Bioactive Aldehydes. J. Biol. Chem. 2008, 283, 21837−21841. (518) Stadtman, E. R. Oxidation of Free Amino Acids and Amino Acid Residues in Proteins by Radiolysis and by Metal-Catalyzed Reactions. Annu. Rev. Biochem. 1993, 62, 797−821. (519) Stadtman, E. R.; Levine, R. L. Free Radical-Mediated Oxidation of Free Amino Acids and Amino Acid Residues in Proteins. Amino Acids 2003, 25, 207−218. (520) Debets, M. F.; van der Doelen, C. W.; Rutjes, F. P.; van Delft, F. L. Azide: A Unique Dipole for Metal-Free Bioorthogonal Ligations. ChemBioChem 2010, 11, 1168−1184. (521) Debets, M. F.; van Berkel, S. S.; Dommerholt, J.; Dirks, A. T.; Rutjes, F. P.; van Delft, F. L. Bioconjugation with Strained Alkenes and Alkynes. Acc. Chem. Res. 2011, 44, 805−815. (522) Sletten, E. M.; Bertozzi, C. R. From Mechanism to Mouse: A Tale of Two Bioorthogonal Reactions. Acc. Chem. Res. 2011, 44, 666− 676. (523) Rostovtsev, V. V.; Green, L. G.; Fokin, V. V.; Sharpless, K. B. A Stepwise Huisgen Cycloaddition Process: Copper(I)-Catalyzed Regioselective ″Ligation″ of Azides and Terminal Alkynes. Angew. Chem., Int. Ed. 2002, 41, 2596−2599. (524) Meldal, M.; Tornoe, C. W. Cu-Catalyzed Azide-Alkyne Cycloaddition. Chem. Rev. 2008, 108, 2952−3015. (525) Kolb, H. C.; Finn, M. G.; Sharpless, K. B. Click Chemistry: Diverse Chemical Function from a Few Good Reactions. Angew. Chem., Int. Ed. 2001, 40, 2004−2021. (526) Wang, Q.; Chan, T. R.; Hilgraf, R.; Fokin, V. V.; Sharpless, K. B.; Finn, M. G. Bioconjugation by Copper(I)-Catalyzed Azide-Alkyne [3 + 2] Cycloaddition. J. Am. Chem. Soc. 2003, 125, 3192−3193. (527) Agard, N. J.; Baskin, J. M.; Prescher, J. A.; Lo, A.; Bertozzi, C. R. A Comparative Study of Bioorthogonal Reactions with Azides. ACS Chem. Biol. 2006, 1, 644−648. (528) del Amo, D. S.; Wang, W.; Jiang, H.; Besanceney, C.; Yan, A. C.; Levy, M.; Liu, Y.; Marlow, F. L.; Wu, P. Biocompatible Copper(I) Catalysts for in Vivo Imaging of Glycans. J. Am. Chem. Soc. 2010, 132, 16893−16899. (529) Spicer, C. D.; Triemer, T.; Davis, B. G. Palladium-Mediated Cell-Surface Labeling. J. Am. Chem. Soc. 2012, 134, 800−803. (530) Li, J.; Lin, S. X.; Wang, J.; Jia, S.; Yang, M. Y.; Hao, Z. Y.; Zhang, X. Y.; Chen, P. R. Ligand-Free Palladium-Mediated SiteSpecific Protein Labeling Inside Gram-Negative Bacterial Pathogens. J. Am. Chem. Soc. 2013, 135, 7330−7338. (531) Saxon, E.; Bertozzi, C. R. Cell Surface Engineering by a Modified Staudinger Reaction. Science 2000, 287, 2007−2010. (532) Blomquist, A. T.; Liu, L. H. Many-Membered Carbon Rings VII. Cyclooctyne. J. Am. Chem. Soc. 1953, 75, 2153−2154.

(533) Wittig, G.; Krebs, A. Zur Existenz Niedergliedriger Cycloalkine I. Chem. Ber. 1961, 94, 3260−3275. (534) Seitz, G.; Pohl, L.; Pohlke, R. 5,6-Didehydro-11,12Dihydrodibenzo[a,E] Cyclooctene. Angew. Chem., Int. Ed. Engl. 1969, 8, 447−448. (535) Agard, N. J.; Prescher, J. A.; Bertozzi, C. R. A Strain-Promoted [3 + 2] Azide-Alkyne Cycloaddition for Covalent Modification of Biomolecules in Living Systems. J. Am. Chem. Soc. 2004, 126, 15046− 15047. (536) Ning, X.; Guo, J.; Wolfert, M. A.; Boons, G. J. Visualizing Metabolically Labeled Glycoconjugates of Living Cells by Copper-Free and Fast Huisgen Cycloadditions. Angew. Chem., Int. Ed. 2008, 47, 2253−2255. (537) Park, M.; Tian, H.; Naganathan, S.; Sakmar, T. P.; Huber, T. Quantitative Multi-color Detection Strategies for Bioorthogonally Labeled GPCRs. Methods Mol. Biol. 2015, 1335, 67−93. (538) Fairbanks, B. D.; Sims, E. A.; Anseth, K. S.; Bowman, C. N. Reaction Rates and Mechanisms for Radical, Photoinitated Addition of Thiols to Alkynes, and Implications for Thiol-Yne Photopolymerizations and Click Reactions. Macromolecules 2010, 43, 4113−4119. (539) van Geel, R.; Pruijn, G. J. M.; van Delft, F. L.; Boelens, W. C. Preventing Thiol-Yne Addition Improves the Specificity of StrainPromoted Azide-Alkyne Cycloaddition. Bioconjugate Chem. 2012, 23, 392−398. (540) Tian, H.; Sakmar, T. P.; Huber, T. A Simple Method for Enhancing the Bioorthogonality of Cyclooctyne Reagent. Chem. Commun. 2016, 52, 5451−5454. (541) Anderton, G. I.; Bangerter, A. S.; Davis, T. C.; Feng, Z. Y.; Furtak, A. J.; Larsen, J. O.; Scroggin, T. L.; Heemstra, J. M. Accelerating Strain-Promoted Azide-Alkyne Cycloaddition Using Micellar Catalysis. Bioconjugate Chem. 2015, 26, 1687−1691. (542) Nguyen, D. P.; Lusic, H.; Neumann, H.; Kapadnis, P. B.; Deiters, A.; Chin, J. W. Genetic Encoding and Labeling of Aliphatic Azides and Alkynes in Recombinant Proteins Via a Pyrrolysyl-tRNA Synthetase/tRNA(CUA) Pair and Click Chemistry. J. Am. Chem. Soc. 2009, 131, 8720−8721. (543) Hancock, S. M.; Uprety, R.; Deiters, A.; Chin, J. W. Expanding the Genetic Code of Yeast for Incorporation of Diverse Unnatural Amino Acids Via a Pyrrolysyl-tRNA Synthetase/tRNA Pair. J. Am. Chem. Soc. 2010, 132, 14819−14824. (544) Lee, Y. J.; Wu, B.; Raymond, J. E.; Zeng, Y.; Fang, X.; Wooley, K. L.; Liu, W. R. A Genetically Encoded Acrylamide Functionality. ACS Chem. Biol. 2013, 8, 1664−1670. (545) Yu, Z. P.; Pan, Y. C.; Wang, Z. Y.; Wang, J. Y.; Lin, Q. Genetically Encoded Cyclopropene Directs Rapid, PhotoclickChemistry-Mediated Protein Labeling in Mammalian Cells. Angew. Chem., Int. Ed. 2012, 51, 10600−10604. (546) Nguyen, D. P.; Elliott, T.; Holt, M.; Muir, T. W.; Chin, J. W. Genetically Encoded 1,2-Aminothiols Facilitate Rapid and Site-Specific Protein Labeling via a Bio-orthogonal Cyanobenzothiazole Condensation. J. Am. Chem. Soc. 2011, 133, 11418−11421. (547) Plass, T.; Milles, S.; Koehler, C.; Schultz, C.; Lemke, E. A. Genetically Encoded Copper-Free Click Chemistry. Angew. Chem., Int. Ed. 2011, 50, 3878−3881. (548) Borrmann, A.; Milles, S.; Plass, T.; Dommerholt, J.; Verkade, J. M.; Wiessler, M.; Schultz, C.; van Hest, J. C.; van Delft, F. L.; Lemke, E. A. Genetic Encoding of a Bicyclo[6.1.0]Nonyne-Charged Amino Acid Enables Fast Cellular Protein Imaging by Metal-Free Ligation. ChemBioChem 2012, 13, 2094−2099. (549) Borrmann, A.; Fatunsin, O.; Dommerholt, J.; Jonker, A. M.; Lowik, D. W. P. M.; van Hest, J. C. M.; van Delft, F. L. StrainPromoted Oxidation-Controlled Cyclooctyne-1,2-Quinone Cycloaddition (SPOCQ) for Fast and Activatable Protein Conjugation. Bioconjugate Chem. 2015, 26, 257−261. (550) Kaya, E.; Vrabel, M.; Deiml, C.; Prill, S.; Fluxa, V. S.; Carell, T. A Genetically Encoded Norbornene Amino Acid for the Mild and Selective Modification of Proteins in a Copper-Free Click Reaction. Angew. Chem., Int. Ed. 2012, 51, 4466−4469. BD

DOI: 10.1021/acs.chemrev.6b00084 Chem. Rev. XXXX, XXX, XXX−XXX

Chemical Reviews

Review

(573) Kuszak, A. J.; Pitchiaya, S.; Anand, J. P.; Mosberg, H. I.; Walter, N. G.; Sunahara, R. K. Purification and Functional Reconstitution of Monomeric μ-Opioid Receptors Allosteric Modulation of Agonist Binding by G(i2). J. Biol. Chem. 2009, 284, 26732−26741. (574) Milligan, G. G Protein-Coupled Receptor Hetero-Dimerization: Contribution to Pharmacology and Function. Br. J. Pharmacol. 2009, 158, 5−14. (575) Briddon, S. J.; Hill, S. J. Pharmacology under the Microscope: The Use of Fluorescence Correlation Spectroscopy to Determine the Properties of Ligand-Receptor Complexes. Trends Pharmacol. Sci. 2007, 28, 637−645. (576) Fricke, F.; Dietz, M. S.; Heilemann, M. Single-Molecule Methods to Study Membrane Receptor Oligomerization. ChemPhysChem 2015, 16, 713−721. (577) Kasai, R. S.; Kusumi, A. Single-Molecule Imaging Revealed Dynamic GPCR Dimerization. Curr. Opin. Cell Biol. 2014, 27, 78−86. (578) Hern, J. A.; Baig, A. H.; Mashanov, G. I.; Birdsall, B.; Corrie, J. E. T.; Lazareno, S.; Molloy, J. E.; Birdsall, N. J. M. Formation and Dissociation of M-1 Muscarinic Receptor Dimers Seen by Total Internal Reflection Fluorescence Imaging of Single Molecules. Proc. Natl. Acad. Sci. U. S. A. 2010, 107, 2693−2698. (579) Kasai, R. S.; Suzuki, K. G. N.; Prossnitz, E. R.; Koyama-Honda, I.; Nakada, C.; Fujiwara, T. K.; Kusumi, A. Full Characterization of GPCR Monomer-Dimer Dynamic Equilibrium by Single Molecule Imaging. J. Cell Biol. 2011, 192, 463−480. (580) Fotiadis, D.; Jastrzebska, B.; Philippsen, A.; Muller, D. J.; Palczewski, K.; Engel, A. Structure of the Rhodopsin Dimer: A Working Model for G Protein-Coupled Receptors. Curr. Opin. Struct. Biol. 2006, 16, 252−259. (581) Endesfelder, U.; Finan, K.; Holden, S. J.; Cook, P. R.; Kapanidis, A. N.; Heilemann, M. Multi-Scale Spatial Organization of Rna Polymerase in Escherichia Coli. Biophys. J. 2013, 105, 172−181. (582) Vobornik, D.; Rouleau, Y.; Haley, J.; Bani-Yaghoub, M.; Taylor, R.; Johnston, L. J.; Pezacki, J. P. Nanoscale Organization of β(2)-Adrenergic Receptor-Venus Fusion Protein Domains on the Surface of Mammalian Cells. Biochem. Biophys. Res. Commun. 2009, 382, 85−90. (583) Annibale, P.; Vanni, S.; Scarselli, M.; Rothlisberger, U.; Radenovic, A. Quantitative Photo Activated Localization Microscopy: Unraveling the Effects of Photoblinking. PLoS One 2011, 6, e22678. (584) Scarselli, M.; Annibale, P.; Radenovic, A. Cell Type-Specific β2-Adrenergic Receptor Clusters Identified Using Photoactivated Localization Microscopy Are Not Lipid Raft Related, but Depend on Actin Cytoskeleton Integrity. J. Biol. Chem. 2012, 287, 16768−16780. (585) Scarselli, M.; Annibale, P.; Gerace, C.; Radenovic, A. Enlightening G Protein-Coupled Receptors on the Plasma Membrane Using Super-Resolution Photoactivated Localization Microscopy. Biochem. Soc. Trans. 2013, 41, 191−196. (586) Lee, S. F.; Vérolet, Q.; Fürstenberg, A. Improved SuperResolution Microscopy with Oxazine Fluorophores in Heavy Water. Angew. Chem., Int. Ed. 2013, 52, 8948−8951. (587) Jonas, K. C.; Fanelli, F.; Huhtaniemi, I. T.; Hanyaloglu, A. C. Single Molecule Analysis of Functionally Asymmetric G Proteincoupled Receptor (GPCR) Oligomers Reveals Diverse Spatial and Structural Assemblies. J. Biol. Chem. 2015, 290, 3875−3892. (588) Truan, Z.; Tarancón Díez, L.; Bönsch, C.; Malkusch, S.; Endesfelder, U.; Munteanu, M.; Hartley, O.; Heilemann, M.; Fürstenberg, A. Quantitative Morphological Analysis of Arrestin2 Clustering Upon G Protein-Coupled Receptor Stimulation by SuperResolution Microscopy. J. Struct. Biol. 2013, 184, 329−334. (589) van Oijen, A. M.; Dixon, N. E. Probing Molecular Choreography through Single-Molecule Biochemistry. Nat. Struct. Mol. Biol. 2015, 22, 948−952. (590) Bockenhauer, S.; Fürstenberg, A.; Xiao Jie, Y.; Kobilka, B. K.; Moerner, W. E. Anti-Brownian ELectrokinetic (ABEL) Trapping of Single β2-Adrenergic Receptors in the Absence and Presence of Agonist. Proc. SPIE 2012, 8228, 822805.

(551) Lang, K.; Davis, L.; Torres-Kolbus, J.; Chou, C.; Deiters, A.; Chin, J. W. Genetically Encoded Norbornene Directs Site-Specific Cellular Protein Labelling Via a Rapid Bioorthogonal Reaction. Nat. Chem. 2012, 4, 298−304. (552) Plass, T.; Milles, S.; Koehler, C.; Szymanski, J.; Mueller, R.; Wiessler, M.; Schultz, C.; Lemke, E. A. Amino Acids for Diels-Alder Reactions in Living Cells. Angew. Chem., Int. Ed. 2012, 51, 4166−4170. (553) Lang, K.; Davis, L.; Wallace, S.; Mahesh, M.; Cox, D. J.; Blackman, M. L.; Fox, J. M.; Chin, J. W. Genetic Encoding of Bicyclononynes and Trans-Cyclooctenes for Site-Specific Protein Labeling in Vitro and in Live Mammalian Cells Via Rapid Fluorogenic Diels-Alder Reactions. J. Am. Chem. Soc. 2012, 134, 10317−10320. (554) Hoyle, C. E.; Bowman, C. N. Thiol-Ene Click Chemistry. Angew. Chem., Int. Ed. 2010, 49, 1540−1573. (555) Jewett, J. C.; Bertozzi, C. R. Cu-Free Click Cycloaddition Reactions in Chemical Biology. Chem. Soc. Rev. 2010, 39, 1272−1279. (556) Nikic, I.; Kang, J. H.; Girona, G. E.; Aramburu, I. V.; Lemke, E. A. Labeling Proteins on Live Mammalian Cells Using Click Chemistry. Nat. Protoc. 2015, 10, 780−791. (557) Le Droumaguet, C.; Wang, C.; Wang, Q. Fluorogenic Click Reaction. Chem. Soc. Rev. 2010, 39, 1233−1239. (558) Grimm, J. B.; Heckman, L. M.; Lavis, L. D. The Chemistry of Small-Molecule Fluorogenic Probes. Prog. Mol. Biol. Transl. Sci. 2013, 113, 1−34. (559) Nadler, A.; Schultz, C. The Power of Fluorogenic Probes. Angew. Chem., Int. Ed. 2013, 52, 2408−2410. (560) Hori, Y.; Kikuchi, K. Protein Labeling with Fluorogenic Probes for No-Wash Live-Cell Imaging of Proteins. Curr. Opin. Chem. Biol. 2013, 17, 644−650. (561) Sivakumar, K.; Xie, F.; Cash, B. M.; Long, S.; Barnhill, H. N.; Wang, Q. A Fluorogenic 1,3-Dipolar Cycloaddition Reaction of 3Azidocoumarins and Acetylenes. Org. Lett. 2004, 6, 4603−4606. (562) Devaraj, N. K.; Weissleder, R. Biomedical Applications of Tetrazine Cycloadditions. Acc. Chem. Res. 2011, 44, 816−827. (563) Devaraj, N. K.; Hilderbrand, S.; Upadhyay, R.; Mazitschek, R.; Weissleder, R. Bioorthogonal Turn-on Probes for Imaging Small Molecules inside Living Cells. Angew. Chem., Int. Ed. 2010, 49, 2869− 2872. (564) Wu, H. X.; Yang, J.; Seckute, J.; Devaraj, N. K. In Situ Synthesis of Alkenyl Tetrazines for Highly Fluorogenic Bioorthogonal Live-Cell Imaging Probes. Angew. Chem., Int. Ed. 2014, 53, 5805−5809. (565) Xie, F.; Sivakumar, K.; Zeng, Q. B.; Bruckman, M. A.; Hodges, B.; Wang, Q. A Fluorogenic ’Click’ Reaction of Azidoanthracene Derivatives. Tetrahedron 2008, 64, 2906−2914. (566) Shieh, P.; Hangauer, M. J.; Bertozzi, C. R. Fluorogenic Azidofluoresceins for Biological Imaging. J. Am. Chem. Soc. 2012, 134, 17428−17431. (567) Shieh, P.; Siegrist, M. S.; Cullen, A. J.; Bertozzi, C. R. Imaging Bacterial Peptidoglycan with Near-Infrared Fluorogenic Azide Probes. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, 5456−5461. (568) Carlson, J. C. T.; Meimetis, L. G.; Hilderbrand, S. A.; Weissleder, R. BODIPY-Tetrazine Derivatives as Superbright Bioorthogonal Turn-on Probes. Angew. Chem., Int. Ed. 2013, 52, 6917− 6920. (569) Sun, X.; Zhang, A.; Baker, B.; Sun, L.; Howard, A.; Buswell, J.; Maurel, D.; Masharina, A.; Johnsson, K.; Noren, C. J.; et al. Development of SNAP-Tag Fluorogenic Probes for Wash-Free Fluorescence Imaging. ChemBioChem 2011, 12, 2217−2226. (570) Lukinavicius, G.; Johnsson, K. Switchable Fluorophores for Protein Labeling in Living Cells. Curr. Opin. Chem. Biol. 2011, 15, 768−774. (571) Fichter, K. M.; Flajolet, M.; Greengard, P.; Vu, T. Q. Kinetics of G Protein-Coupled Receptor Endosomal Trafficking Pathways Revealed by Single Quantum Dots. Proc. Natl. Acad. Sci. U. S. A. 2010, 107, 18658−18663. (572) McLean, A. J.; Bevan, N.; Rees, S.; Milligan, G. Visualizing Differences in Ligand Regulation of Wild-Type and Constitutively Active Mutant β(2)-Adrenoceptor-Green Fluorescent Protein Fusion Proteins. Mol. Pharmacol. 1999, 56, 1182−1191. BE

DOI: 10.1021/acs.chemrev.6b00084 Chem. Rev. XXXX, XXX, XXX−XXX

Chemical Reviews

Review

(591) Hwang, H.; Myong, S. Protein Induced Fluorescence Enhancement (PIFE) for Probing Protein-Nucleic Acid Interactions. Chem. Soc. Rev. 2014, 43, 1221−1229. (592) Vafabakhsh, R.; Levitz, J.; Isacoff, E. Y. Conformational Dynamics of a Class C G Protein-Coupled Receptor. Nature 2015, 524, 497−501. (593) Rose, R. H.; Briddon, S. J.; Hill, S. J. A Novel Fluorescent Histamine H1 Receptor Antagonist Demonstrates the Advantage of Using Fluorescence Correlation Spectroscopy to Study the Binding of Lipophilic Ligands. Br. J. Pharmacol. 2012, 165, 1789−1800. (594) Hegener, O.; Prenner, L.; Runkel, F.; Baader, S. L.; Kappler, J.; Häberlein, H. Dynamics of β2-Adrenergic Receptor−Ligand Complexes on Living Cells. Biochemistry 2004, 43, 6190−6199. (595) Russel, D.; Lasker, K.; Phillips, J.; Schneidman-Duhovny, D.; Velazquez-Muriel, J. A.; Sali, A. The Structural Dynamics of Macromolecular Processes. Curr. Opin. Cell Biol. 2009, 21, 97−108. (596) Zocher, M.; Bippes, C. A.; Zhang, C.; Mueller, D. J. SingleMolecule Force Spectroscopy of G Protein-Coupled Receptors. Chem. Soc. Rev. 2013, 42, 7801−7815. (597) Nogales, E. The Development of Cryo-EM into a Mainstream Structural Biology Technique. Nat. Methods 2015, 13, 24−27. (598) Westfield, G. H.; Rasmussen, S. G. F.; Su, M.; Dutta, S.; DeVree, B. T.; Chung, K. Y.; Calinski, D.; Velez-Ruiz, G.; Oleskie, A. N.; Pardon, E.; et al. Structural Flexibility of the Gαs α-Helical Domain in the β(2)-Adrenoceptor Gs Complex. Proc. Natl. Acad. Sci. U. S. A. 2011, 108, 16086−16091. (599) Shukla, A. K.; Westfield, G. H.; Xiao, K.; Reis, R. I.; Huang, L.Y.; Tripathi-Shukla, P.; Qian, J.; Li, S.; Blanc, A.; Oleskie, A. N.; et al. Visualization of Arrestin Recruitment by a G Protein-Coupled Receptor. Nature 2014, 512, 218−222. (600) Yang, L.; Yang, D.; de Graaf, C.; Moeller, A.; West, G. M.; Dharmarajan, V.; Wang, C.; Siu, F. Y.; Song, G.; Reedtz-Runge, S. Conformational States of the Full-Length Glucagon Receptor. Nat. Commun. 2015, 6, 7859. (601) Panneels, V.; Wu, W.; Tsai, C.-J.; Nogly, P.; Rheinberger, J.; Jaeger, K.; Cicchetti, G.; Gati, C.; Kick, L. M.; Sala, L.; et al. TimeResolved Structural Studies with Serial Crystallography: A New Light on Retinal Proteins. Struct. Dyn. 2015, 2, 041718. (602) Zhang, H.; Unal, H.; Gati, C.; Han, G. W.; Liu, W.; Zatsepin, N. A.; James, D.; Wang, D.; Nelson, G.; Weierstall, U.; et al. Structure of the Angiotensin Receptor Revealed by Serial Femtosecond Crystallography. Cell 2015, 161, 833−844. (603) Dror, R. O.; Arlow, D. H.; Maragakis, P.; Mildorf, T. J.; Pan, A. C.; Xu, H.; Borhani, D. W.; Shaw, D. E. Activation Mechanism of the β(2)-Adrenergic Receptor. Proc. Natl. Acad. Sci. U. S. A. 2011, 108, 18684−18689. (604) Dror, R. O.; Mildorf, T. J.; Hilger, D.; Manglik, A.; Borhani, D. W.; Arlow, D. H.; Philippsen, A.; Villanueva, N.; Yang, Z.; Lerch, M. T.; et al. Structural Basis for Nucleotide Exchange in Heterotrimeric G Proteins. Science 2015, 348, 1361−1365. (605) Dror, R. O.; Dirks, R. M.; Grossman, J. P.; Xu, H.; Shaw, D. E. Biomolecular Simulation: A Computational Microscope for Molecular Biology. Annu. Rev. Biophys. 2012, 41, 429−452. (606) Liebmann, T.; Kruusmagi, M.; Sourial-Bassillious, N.; Bondar, A.; Svenningsson, P.; Flajolet, M.; Greengard, P.; Scott, L.; Brismar, H.; Aperia, A. A Noncanonical Postsynaptic Transport Route for a GPCR Belonging to the Serotonin Receptor Family. J. Neurosci. 2012, 32, 17998−18008. (607) Herrick-Davis, K.; Grinde, E.; Cowan, A.; Mazurkiewicz, J. E. Fluorescence Correlation Spectroscopy Analysis of Serotonin, Adrenergic, Muscarinic, and Dopamine Receptor Dimerization: The Oligomer Number Puzzle. Mol. Pharmacol. 2013, 84, 630−642. (608) Liu, W.; Wacker, D.; Gati, C.; Han, G. W.; James, D.; Wang, D.; Nelson, G.; Weierstall, U.; Katritch, V.; Barty, A.; et al. Serial Femtosecond Crystallography of G Protein-Coupled Receptors. Science 2013, 342, 1521−1524. (609) Herrick-Davis, K.; Grinde, E.; Lindsley, T.; Cowan, A.; Mazurkiewicz, J. E. Oligomer Size of the Serotonin 5-Hydroxytryptamine 2C (5-HT2C) Receptor Revealed by Fluorescence Correlation

Spectroscopy with Photon Counting Histogram Analysis: Evidence for Homodimers without Monomers or Tetramers. J. Biol. Chem. 2012, 287, 23604−23614. (610) Briddon, S. J.; Middleton, R. J.; Yates, A. S.; George, M. W.; Kellam, B.; Hill, S. J. Application of Fluorescence Correlation Spectroscopy to the Measurement of Agonist Binding to a G Protein-Coupled Receptor at the Single Cell Level. Faraday Discuss. 2004, 126, 197−207. (611) Briddon, S. J.; Gandia, J.; Amaral, O. B.; Ferre, S.; Lluis, C.; Franco, R.; Hill, S. J.; Ciruela, F. Plasma Membrane Diffusion of G Protein-Coupled Receptor Oligomers. Biochim. Biophys. Acta, Mol. Cell Res. 2008, 1783, 2262−2268. (612) Middleton, R. J.; Briddon, S. J.; Cordeaux, Y.; Yates, A. S.; Dale, C. L.; George, M. W.; Baker, J. G.; Hill, S. J.; Kellam, B. New Fluorescent Adenosine a(1)-Receptor Agonists That Allow Quantification of Ligand-Receptor Interactions in Microdomains of Single Living Cells. J. Med. Chem. 2007, 50, 782−793. (613) Keuerleber, S.; Thurner, P.; Gruber, C. W.; Zezula, J.; Freissmuth, M. Reengineering the Collision Coupling and Diffusion Mode of the A(2a)-Adenosine Receptor: Palmitoylation in Helix 8 Relieves Confinement. J. Biol. Chem. 2012, 287, 42104−42118. (614) Thurner, P.; Gsandtner, I.; Kudlacek, O.; Choquet, D.; Nanoff, C.; Freissmuth, M.; Zezula, J. A Two-State Model for the Diffusion of the A(2a) Adenosine Receptor in Hippocampal Neurons. J. Biol. Chem. 2014, 289, 9263−9274. (615) Cordeaux, Y.; Briddon, S. J.; Alexander, S. P.; Kellam, B.; Hill, S. J. Agonist-Occupied A3 Adenosine Receptors Exist within Heterogeneous Complexes in Membrane Microdomains of Individual Living Cells. FASEB J. 2007, 22, 850−860. (616) Corriden, R.; Kilpatrick, L. E.; Kellam, B.; Briddon, S. J.; Hill, S. J. Kinetic Analysis of Antagonist-Occupied Adenosine-A(3) Receptors within Membrane Microdomains of Individual Cells Provides Evidence of Receptor Dimerization and Allosterism. FASEB J. 2014, 28, 4211−4222. (617) Perez, J.-B.; Segura, J.-M.; Abankwa, D.; Piguet, J.; Martinez, K. L.; Vogel, H. Monitoring the Diffusion of Single Heterotrimeric G Proteins in Supported Cell-Membrane Sheets Reveals Their Partitioning into Microdomains. J. Mol. Biol. 2006, 363, 918−930. (618) Wagner, J.; Sungkaworn, T.; Heinze, K. G.; Lohse, M. J.; Calebiro, D. Single-Molecule Fluorescence Microscopy for the Analysis of Fast Receptor Dynamics. Methods Mol. Biol. 2015, 1335, 53−66. (619) Zocher, M.; Roos, C.; Wegmann, S.; Bosshart, P. D.; Doetsch, V.; Bernhard, F.; Mueller, D. J. Single-Molecule Force Spectroscopy from Nanodiscs: An Assay to Quantify Folding, Stability, and Interactions of Native Membrane Proteins. ACS Nano 2012, 6, 961−971. (620) Müller, D. J.; Kessler, M.; Oesterhelt, F.; Mö ller, C.; Oesterhelt, D.; Gaub, H. Stability of Bacteriorhodopsin α-Helices and Loops Analyzed by Single-Molecule Force Spectroscopy. Biophys. J. 2002, 83, 3578−3588. (621) Sapra, K. T.; Park, P. S. H.; Palczewski, K.; Muller, D. J. Mechanical Properties of Bovine Rhodopsin and Bacteriorhodopsin: Possible Roles in Folding and Function†. Langmuir 2008, 24, 1330− 1337. (622) Sapra, K. T.; Balasubramanian, G. P.; Labudde, D.; Bowie, J. U.; Muller, D. J. Point Mutations in Membrane Proteins Reshape Energy Landscape and Populate Different Unfolding Pathways. J. Mol. Biol. 2008, 376, 1076−1090. (623) Sapra, K. T.; Doehner, J.; Renugopalakrishnan, V.; Padrós, E.; Muller, D. J. Role of Extracellular Glutamic Acids in the Stability and Energy Landscape of Bacteriorhodopsin. Biophys. J. 2008, 95, 3407− 3418. (624) Sapra, K. T.; Besir, H.; Oesterhelt, D.; Muller, D. J. Characterizing Molecular Interactions in Different Bacteriorhodopsin Assemblies by Single-molecule Force Spectroscopy. J. Mol. Biol. 2006, 355, 640−650. (625) Philip, F.; Sengupta, P.; Scarlata, S. Signaling through a G Protein-Coupled Receptor and Its Corresponding G Protein Follows a BF

DOI: 10.1021/acs.chemrev.6b00084 Chem. Rev. XXXX, XXX, XXX−XXX

Chemical Reviews

Review

Stoichiometrically Limited Model. J. Biol. Chem. 2007, 282, 19203− 19216. (626) Valentine, C. D.; Haggie, P. M. Confinement of β(1)- and β(2)-Adrenergic Receptors in the Plasma Membrane of Cardiomyocyte-Like H9c2 Cells Is Mediated by Selective Interactions with PDZ Domain and A-Kinase Anchoring Proteins but Not Caveolae. Mol. Biol. Cell 2011, 22, 2970−2982. (627) Peisley, A.; Skiniotis, G. 2D Projection Analysis of GPCR Complexes by Negative Stain Electron Microscopy. Methods Mol. Biol. 2015, 1335, 29−38. (628) Sungkaworn, T.; Rieken, F.; Lohse, M. J.; Calebiro, D. HighResolution Spatiotemporal Analysis of Receptor Dynamics by SingleMolecule Fluorescence Microscopy. J. Vis. Exp. 2014, DOI: 10.3791/ 51784. (629) Zocher, M.; Fung, J. J.; Kobilka, B. K.; Mueller, D. J. LigandSpecific Interactions Modulate Kinetic, Energetic, and Mechanical Properties of the Human β(2) Adrenergic Receptor. Structure 2012, 20, 1391−1402. (630) Zocher, M.; Zhang, C.; Rasmussen, S. G. F.; Kobilka, B. K.; Mueller, D. J. Cholesterol Increases Kinetic, Energetic, and Mechanical Stability of the Human β(2)-Adrenergic Receptor. Proc. Natl. Acad. Sci. U. S. A. 2012, 109, E3463−E3472. (631) Licht, S. S.; Sonnleitner, A.; Weiss, S.; Schultz, P. G. A Rugged Energy Landscape Mechanism for Trapping of Transmembrane Receptors During Endocytosis. Biochemistry 2003, 42, 2916−2925. (632) Sergé, A.; de Keijzer, S.; Van Hemert, F.; Hickman, M. R.; Hereld, D.; Spaink, H. P.; Schmidt, T.; Snaar-Jagalska, B. E. Quantification of GPCR Internalization by Single-Molecule Microscopy in Living Cells. Integr. Biol. 2011, 3, 675−683. (633) Mikasova, L.; Groc, L.; Choquet, D.; Manzoni, O. J. Altered Surface Trafficking of Presynaptic Cannabinoid Type 1 Receptor in and out Synaptic Terminals Parallels Receptor Desensitization. Proc. Natl. Acad. Sci. U. S. A. 2008, 105, 18596−18601. (634) Pramanik, A.; Olsson, M.; Langel, U.; Bartfai, T.; Rigler, R. Fluorescence Correlation Spectroscopy Detects Galanin Receptor Diversity on Insulinoma Cells. Biochemistry 2001, 40, 10839−10845. (635) Sergé, A.; Fourgeaud, L.; Hemar, A.; Choquet, D. Receptor Activation and Homer Differentially Control the Lateral Mobility of Metabotropic Glutamate Receptor 5 in the Neuronal Membrane. J. Neurosci. 2002, 22, 3910−3920. (636) Sergé, A.; Fourgeaud, L.; Hémar, A.; Choquet, D. Active Surface Transport of Metabotropic Glutamate Receptors through Binding to Microtubules and Actin Flow. J. Cell Sci. 2003, 116, 5015− 5022. (637) Smith, S. M.; Lei, Y.; Liu, J.; Cahill, M. E.; Hagen, G. M.; Barisas, B. G.; Roess, D. A. Luteinizing Hormone Receptors Translocate to Plasma Membrane Microdomains after Binding of Human Chorionic Gonadotropin. Endocrinology 2006, 147, 1789− 1795. (638) Lill, Y.; Martinez, K. L.; Lill, M. A.; Meyer, B. H.; Vogel, H.; Hecht, B. Kinetics of the Initial Steps of G Protein-Coupled ReceptorMediated Cellular Signaling Revealed by Single-Molecule Imaging. ChemPhysChem 2005, 6, 1633−1640. (639) Prummer, M.; Meyer, B. H.; Franzini, R.; Segura, J. M.; George, N.; Johnsson, K.; Vogel, H. Post-Translational Covalent Labeling Reveals Heterogeneous Mobility of Individual G ProteinCoupled Receptors in Living Cells. ChemBioChem 2006, 7, 908−911. (640) Kilpatrick, L. E.; Briddon, S. J.; Holliday, N. D. Fluorescence Correlation Spectroscopy, Combined with Bimolecular Fluorescence Complementation, Reveals the Effects of β-Arrestin Complexes and Endocytic Targeting on the Membrane Mobility of Neuropeptide Y Receptors. Biochim. Biophys. Acta, Mol. Cell Res. 2012, 1823, 1068− 1081. (641) Daumas, F.; Destainville, N.; Millot, C.; Lopez, A.; Dean, D.; Salome, L. Confined Diffusion without Fences of a G Protein-Coupled Receptor as Revealed by Single Particle Tracking. Biophys. J. 2003, 84, 356−366. (642) Daumas, F.; Destainville, N.; Millot, C.; Lopez, A.; Dean, D.; Salome, L. Interprotein Interactions Are Responsible for the Confined

Diffusion of a G Protein-Coupled Receptor at the Cell Surface. Biochem. Soc. Trans. 2003, 31, 1001−1005. (643) Suzuki, K.; Ritchie, K.; Kajikawa, E.; Fujiwara, T.; Kusumi, A. Rapid Hop Diffusion of a G Protein-Coupled Receptor in the Plasma Membrane as Revealed by Single-Molecule Techniques. Biophys. J. 2005, 88, 3659−3680. (644) Leutenegger, M.; Lasser, T.; Sinner, E.-K.; Robelek, R. Imaging of G Protein-Coupled Receptors in Solid-Supported Planar Lipid Membranes. Biointerphases 2008, 3, FA136−FA145. (645) Märki, I.; Leutenegger, M.; Geissbuehler, M.; Robelek, R.; Sinner, E.-K.; Lasser, T. Imaging of G Protein-Coupled Receptors in Solid-Supported Planar Membranes at the Single Molecule Level. Proc. SPIE 2008, 6862. (646) Jacquier, V.; Prummer, M.; Segura, J. M.; Pick, H.; Vogel, H. Visualizing Odorant Receptor Trafficking in Living Cells Down to the Single-Molecule Level. Proc. Natl. Acad. Sci. U. S. A. 2006, 103, 14325−14330. (647) Alsteens, D.; Pfreundschuh, M.; Zhang, C.; Spoerri, P. M.; Coughlin, S. R.; Kobilka, B. K.; Muller, D. J. Imaging G ProteinCoupled Receptors While Quantifying Their Ligand-Binding FreeEnergy Landscape. Nat. Methods 2015, 12, 845. (648) Pfreundschuh, M.; Alsteens, D.; Wieneke, R.; Zhang, C.; Coughlin, S. R.; Tampe, R.; Kobilka, B. K.; Muller, D. J. Identifying and Quantifying Two Ligand-Binding Sites While Imaging Native Human Membrane Receptors by AFM. Nat. Commun. 2015, 6, 8857. (649) Zelman-Femiak, M.; Wang, K.; Gromova, K. V.; Knaus, P.; Harms, G. S. Covalent Quantum Dot Receptor Linkage Via the Acyl Carrier Protein for Single-Molecule Tracking, Internalization, and Trafficking Studies. BioTechniques 2010, 49, 574−579. (650) Alexiev, U.; Farrens, D. L. Fluorescence Spectroscopy of Rhodopsins: Insights and Approaches. Biochim. Biophys. Acta, Bioenerg. 2014, 1837, 694−709. (651) Kim, T.-Y.; Uji-i, H.; Moeller, M.; Muls, B.; Hofkens, J.; Alexiev, U. Monitoring the Interaction of a Single G Protein Key Binding Site with Rhodopsin Disk Membranes upon Light Activation. Biochemistry 2009, 48, 3801−3803. (652) Kim, T.-Y.; Schlieter, T.; Haase, S.; Alexiev, U. Activation and Molecular Recognition of the GPCR Rhodopsin - Insights from TimeResolved Fluorescence Depolarisation and Single Molecule Experiments. Eur. J. Cell Biol. 2012, 91, 300−310. (653) Kirchberg, K.; Kim, T.-Y.; Haase, S.; Alexiev, U. Functional Interaction Structures of the Photochromic Retinal Protein Rhodopsin. Photochem. Photobiol. Sci. 2010, 9, 226−233. (654) Sapra, K. T.; Park, P. S. H.; Filipek, S.; Engel, A.; Muller, D. J.; Palczewski, K. Detecting Molecular Interactions That Stabilize Native Bovine Rhodopsin. J. Mol. Biol. 2006, 358, 255−269. (655) Kawamura, S.; Colozo, A. T.; Müller, D. J.; Park, P. S. H. Conservation of Molecular Interactions Stabilizing Bovine and Mouse Rhodopsin. Biochemistry 2010, 49, 10412−10420. (656) Kawamura, S.; Colozo, A. T.; Ge, L.; Muller, D. J.; Park, P. S. Structural, Energetic, and Mechanical Perturbations in Rhodopsin Mutant That Causes Congenital Stationary Night Blindness. J. Biol. Chem. 2012, 287, 21826−21835. (657) Park, P. S.; Sapra, K. T.; Kolinski, M.; Filipek, S.; Palczewski, K.; Muller, D. J. Stabilizing Effect of Zn2+ in Native Bovine Rhodopsin. J. Biol. Chem. 2007, 282, 11377−11385. (658) Park, P. S. H.; Sapra, K. T.; Jastrzebska, B.; Maeda, T.; Maeda, A.; Pulawski, W.; Kono, M.; Lem, J.; Crouch, R. K.; Filipek, S.; et al. Modulation of Molecular Interactions and Function by Rhodopsin Palmitylation. Biochemistry 2009, 48, 4294−4304. (659) Kawamura, S.; Gerstung, M.; Colozo, Alejandro T.; Helenius, J.; Maeda, A.; Beerenwinkel, N.; Park, Paul S. H.; Müller, Daniel J. Kinetic, Energetic, and Mechanical Differences between Dark-State Rhodopsin and Opsin. Structure 2013, 21, 426−437. (660) Jastrzebska, B.; Ringler, P.; Palczewski, K.; Engel, A. The Rhodopsin-Transducin Complex Houses Two Distinct Rhodopsin Molecules. J. Struct. Biol. 2013, 182, 164−172. (661) Cisneros, D. A.; Oberbarnscheidt, L.; Pannier, A.; Klare, J. P.; Helenius, J.; Engelhard, M.; Oesterhelt, F.; Muller, D. J. Transducer BG

DOI: 10.1021/acs.chemrev.6b00084 Chem. Rev. XXXX, XXX, XXX−XXX

Chemical Reviews

Review

Binding Establishes Localized Interactions to Tune Sensory Rhodopsin II. Structure 2008, 16, 1206−1213. (662) Milenkovic, L.; Weiss, L. E.; Yoon, J.; Roth, T. L.; Su, Y. S.; Sahl, S. J.; Scott, M. P.; Moerner, W. E. Single-Molecule Imaging of Hedgehog Pathway Protein Smoothened in Primary Cilia Reveals Binding Events Regulated by Patched1. Proc. Natl. Acad. Sci. U. S. A. 2015, 112, 8320−8325. (663) Ye, F.; Breslow, D. K.; Koslover, E. F.; Spakowitz, A. J.; Nelson, W. J.; Nachury, M. V. Single Molecule Imaging Reveals a Major Role for Diffusion in the Exploration of Ciliary Space by Signaling Receptors. eLife 2013, 2, e00654. (664) Patel, R. C.; Kumar, U.; Lamb, D. C.; Eid, J. S.; Rocheville, M.; Grant, M.; Rani, A.; Hazlett, T.; Patel, S. C.; Gratton, E.; et al. Ligand Binding to Somatostatin Receptors Induces Receptor-Specific Oligomer Formation in Live Cells. Proc. Natl. Acad. Sci. U. S. A. 2002, 99, 3294−3299. (665) Dougherty, D. A. Unnatural Amino Acids as Probes of Protein Structure and Function. Curr. Opin. Chem. Biol. 2000, 4, 645−652. (666) Daggett, K. A.; Sakmar, T. P. Site-Specific in Vitro and in Vivo Incorporation of Molecular Probes to Study G Protein-Coupled Receptor. Curr. Opin. Chem. Biol. 2011, 15, 392−398. (667) Meimetis, L. G.; Carlson, J. C.; Giedt, R. J.; Kohler, R. H.; Weissleder, R. Ultrafluorogenic Coumarin-Tetrazine Probes for RealTime Biological Imaging. Angew. Chem., Int. Ed. 2014, 53, 7531−7534. (668) Harris, L. J.; Larson, S. B.; Hasel, K. W.; McPherson, A. Refined structure of an intact IgG2a monoclonal antibody. Biochemistry 1997, 36, 1581−1597. (669) Yang, F.; Moss, L. G.; Phillips, G. N., Jr. The Molecular Structure of Green Fluorescent Protein. Nat. Biotechnol. 1996, 14, 1246−1251. (670) Loening, A. M.; Fenn, T. D.; Gambhir, S. S. Crystal Structures of the Luciferase and Green Fluorescent Protein from Renilla Reniformis. J. Mol. Biol. 2007, 374, 1017−1028. (671) Humphrey, W.; Dalke, A.; Schulten, K. VMD: Visual Molecular Dynamics. J. Mol. Graphics 1996, 14, 33−38. (672) Smith, A. M.; Nie, S. Semiconductor Nanocrystals: Structure, Properties, and Band Gap Engineering. Acc. Chem. Res. 2010, 43, 190− 200.

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DOI: 10.1021/acs.chemrev.6b00084 Chem. Rev. XXXX, XXX, XXX−XXX