Review
pubs.acs.org/CR
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
B
B
C
C
C
D
F
F
F
G
G
H
H
H
I
J
L
L
L
L
M
M
M
M
N
O
P
P
Q
Q
Q
Q
Q
R
S
S
S
S
S
S
S
T
T
Special Issue: G-Protein Coupled Receptors
N
Received: February 1, 2016
A
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
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.
U
W
W
W
X
Z
AE
AE
AE
AE
AF
AF
AF
AG
AM
AN
AO
AO
AP
AP
AP
AP
AP
AP
AP
AP
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
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
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX
Chemical Reviews
Review
(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.
BH
DOI: 10.1021/acs.chemrev.6b00084
Chem. Rev. XXXX, XXX, XXX−XXX