The Growing and Glowing Toolbox of Fluorescent and Photoactive Proteins

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Series: Superlative Sequels
Feature Review
The Growing and Glowing
Toolbox of Fluorescent and
Photoactive Proteins
Erik A. Rodriguez,
1,
* Robert E. Campbell,
2,
* John Y. Lin,
3,
*
Michael Z. Lin,
4,5,
* Atsushi Miyawaki,
6,
* Amy E. Palmer,
7,
*
Xiaokun Shu,
8,9,
* Jin Zhang,
1,*
and Roger Y. Tsien
1,10,
*
Over the past 20 years, protein engineering has been extensively used to
improve and modify the fundamental properties of fluorescent proteins (FPs)
with the goal of adapting them for a fantastic range of applications. FPs have
been modified by a combination of rational design, structure-based mutagene-
sis, and countless cycles of directed evolution (gene diversification followed by
selection of clones with desired properties) that have collectively pushed the
properties to photophysical and biochemical extremes. In this review, we
provide both a summary of the progress that has been made during the past
two decades, and a broad overview of the current state of FP development and
applications in mammalian systems.
Prototypical FPs
The initial demonstration in 1994 that the Aequorea victoria jellyfish (class Hydrozoa) green
fluorescent protein (avGFP; Figure 1A) could function as a genetically encodable fluorescent tag
[1] was followed by rapid-fire protein-engineering efforts to fine tune its biochemical and
fluorescent properties and expand the color palette to encompass blue, cyan, and yellow
variants [2,3]. Further genomic exploration of marine organisms soon led to the discovery of
additional prototypical FPs (i.e., homologs of avGFP) from class Anthozoa [4], including the
Discosoma striata mushroom anemone, which gave rise to DsRed and subsequent mFruit
progeny (Figure 1B) [5,6], and the sea anemone Entacmaea quadricolor, which yielded eqFP,
TagRFP, mKate, and mRuby derivatives, among others [7–10]. FPs from these species
extended the color palette into the orange, red, and [15_TD$DIFF]farred. Accompanying this evolution of
prototypical FPs, new classes of nonprototypical FP based on the binding of flavin mononucle-
otide (FMN) [11], phycocyanobilin [12], biliverdin [13], or bilirubin [14], have emerged in recent
years. Figure 2A shows the trajectory of FP development, with emphasis on the wild-type roots
of modern variants.
One of the primary goals of FP engineering has been to develop brighter FPs. The brighter an FP,
the lower the intracellular concentration that can be reliably imaged with sufficient fluorescent
contrast relative to the autofluorescent background. The intrinsic fluorescent brightness (see
Glossary)ofafluorophore is proportional to the product of the molecular extinction coefficient (e)
and the fluorescence quantum yield (F). However, practically speaking, the apparent brightness
in cells is highly dependent on additional environmental factors. Therefore, many mutational
strategies have targeted improving gene expression by optimizing codon usage, facilitating
protein folding, promoting the extent and rate of chromophore maturation, and increasing
Trends
Monomeric red and far-red FPs and
indicators now perform nearly as well
as the best green FPs (and indicators).
Reversible and irreversible photochro-
mism in FPs can be exploited to
increase optical resolution and improve
contrast compared with traditional
fluorescence microscopy.
Infrared FPs (IFPs) are becoming ever
more useful as labels for various pro-
teins that allow correct localization and
whole-animal imaging. IFPs can serve
as an additional fluorescent ‘color’for
simultaneous imaging with visible FP-
labeled proteins.
Bacterial phytochrome (BphP)-based
IFPs provide a new scaffold for engi-
neering fluorogenic indicators, which
are ideal to visualize spatiotemporal
dynamics of cell signaling in vivo.
Small ultra-red FP ([3_TD$DIFF]smURFP) is the
brightest far-red nonprototypical FP
(comparable with EGFP) and is extre-
mely photostable. [3_TD$DIFF]smURFP may prove
particularly usefulas a photostable FPfor
super-resolution imaging and as a FRET
acceptor for biosensing applications.
The engineering of new fluorescent
indicators that combine features of
prototypical FP-based indicators with
photochromic proteins can reveal the
cellular maps of biochemical activities
in super-resolution.
FPs can be used as optogenetic actua-
tors to manipulate cellular and protein
functions through chromophore-
assisted light inactivation or light-con-
trolled protein oligomerization.
Trends in Biochemical Sciences, February 2017, Vol. 42, No. 2 http://dx.doi.org/10.1016/j.tibs.2016.09.010 111
© 2016 Elsevier Ltd. All rights reserved.

protein stability by reducing the influence of environmental perturbations, such as pH and O
2
.
Given that most FPs are naturally dimeric or tetrameric, substantial effort has also been aimed at
engineering monomeric variants [5].Figure 2B provides a summary of FP brightness with
respect to the wavelength of maximum excitation and with reference to commonly available
laser lines.
Despite 20 years of FP engineering, there remains substantial room for improvement of FPs[16_TD$DIFF] (see
Outstanding Questions). For instance, two new GFPs, mClover3 [15] and mNeonGreen [16],
have recently emerged as the new benchmark for brightness (approximately a 2.5-fold increase
compared with mEGFP). mClover3 was engineered from avGFP by combining key insights
accumulated from years of evolution, including the identification of positions that influenced the
H-bonding network surrounding the chromophore, stacking residues that shift the energy of
emission, and residues that influence photostability and folding [15,17]. By contrast, mNeon-
Green emerged from an underexplored lineage and was created by incorporating 21 mutations
into the yellow FP derived from Branchiostoma lanceolatum, LanYFP [16], suggesting that there
remains untapped potential in additional marine species. These examples reveal that there may
be considerable room for improvement of prototypical FPs using approaches such as the meta-
analysis of protein-engineering efforts and identifying new lineages of b-barrel FPs. Yet another
approach is to discover or engineer nonprototypical classes of FP, such as FMN-binding
proteins, which have the advantage (relative to avGFP homologs) of developing fluorescence
in low-oxygen environments [11,18,19].
1
Department of Pharmacology,
University of California, San Diego, La
Jolla, CA 92093, USA
2
Department of Chemistry, University
of Alberta, Edmonton, AB, T6G 2G2,
Canada
3
School of Medicine, University of
Tasmania, Hobart, TAS 7000, Australia
4
Department of Bioengineering,
Stanford University, Stanford, CA,
94305, USA
5
Department of Pediatrics, Stanford
University, Stanford, CA, 94305, USA
6
Laboratory for Cell Function
Dynamics, Brain Science Institute,
RIKEN, 2-1 Hirosawa, Wako, Saitama,
351-0198, Japan
7
Department of Chemistry and
Biochemistry, BioFrontiers Institute,
University of Colorado, Boulder, CO,
80303, USA
8
Department of Pharmaceutical
Chemistry, University of California,
San Francisco, San Francisco, CA,
94158, USA
9
Cardiovascular Research Institute,
University of California, San Francisco,
San Francisco, CA, 94158, USA
10
Howard Hughes Medical Institute,
University [10_TD$DIFF]of California, [11_TD$DIFF]San Diego, La
Jolla, CA,[12_TD$DIFF] 92093, USA
*Correspondence: ear001@ucsd.edu
(E.A. Rodriguez), rc4@ualberta.ca
(R.E. Campbell), john.lin@utas.edu.au
(J.Y. Lin), mzlin@stanford.edu (M.Z. Lin),
matsushi@brain.riken.go.jp
(A. Miyawaki),
amy.palmer@colorado.edu
(A.E. Palmer), xiaokun.shu@ucsf.edu
(X. Shu), jzhang32@ucsd.edu (J. Zhang),
rtsien@ucsd.edu (R.Y. Tsien).
(A)
(C) (D) (E)
(B)
Figure 1. Structural Representations of Fluorescent Proteins (FPs). (A) Aequorea green FP (GFP) with the
chromophore represented in green [Protein Data Bank (PDB) ID 1EMA) [160]. (B) The monomeric DsRed-derived RFP
known as mCherry with the chromophore represented in orange (PDB ID 2H5Q) [161]. (C) IFP2.0 derived from a
bacteriophytochrome with bound biliverdin in purple (PDB ID 4CQH) [32]. (D) A representation of a cyanobacterial
allophycocyanin /-subunit that is homologous to small ultra-red FP ([3_TD$DIFF]smURFP), with bound biliverdin shown in red
(PDB 4PO5) [162]. (E) Structure of UnaG with bound bilirubin shown in green (PDB ID 4I3B) [14].
112 Trends in Biochemical Sciences, February 2017, Vol. 42, No. 2

Glossary
Intensiometric fluorescent signal:
a change in fluorescence intensity (i.
e., either an increase or decrease) at
a single wavelength.
Intrinsic fluorescent brightness:
the product of the molecular
extinction coefficient (e) and the
fluorescence quantum yield (F)
Optogenetic actuator: a genetically
encoded protein that undergoes an
illumination-dependent change in
function that induces, disrupts, or
otherwise changes a cellular function.
Photoactivated localization
microscopy (PALM) and
stochastic optical reconstruction
microscopy (STORM): super-
resolution imaging modality in which
an image is constructed from a
multitude of single fluorophore
localizations, where each localization
is determined to a resolution higher
than the diffraction limit.
Ratiometric fluorescent signal: a
change in the ratio of fluorescence
intensity at one wavelength relative to
the fluorescent intensity at a second
wavelength.
Stimulated emission depletion
(STED): a super-resolution imaging
modality in which a first laser beam is
used to excite fluorescence and a
second donut-shaped laser beam is
used to confine the size of the
excitation spot to smaller than the
diffraction limit.
1994
1995
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2005
2006
2007
2008
2009
2010
2011
2012
2013
2014
2015
450 500 550 600 650 700
Emission wavelength (nm)
3
2.5
2
1.5
1
0.5
0
350 400 450 500
Excitaon wavelength (nm)
Brightness (ε × Φ) Relave to EGFP
550 600 650 700
(A)
(B)
Yea r
Figure 2. An Overview of Genetically Encoded Fluorophores Introduced since the Advent of Wild-Type
Aequorea victoria Green Fluorescent Protein (avGFP) in 1994. (A) This chart includes fluorescent proteins (FPs) that
are 11-stranded b-barrel homologs of avGFP, as well as proteins that are fluorescent when bound to a biliverdin or bilirubin
chromophore. Placement along the vertical axis represents the year in which the FP was first introduced, while placement
along the horizontal axis represents the peak emission wavelength. (B) Intrinsic fluorescent brightness (eF) of selected
FPs is plotted relativ e to EGFP as a function of peak excitation wavelength. Vertical lines at 405 nm, 440 nm, 488 nm,
515 nm, 561 nm, 591 nm, and 647 nm correspond to commonly available laser lines on fluorescence microscopes.
Trends in Biochemical Sciences, February 2017, Vol. 42, No. 2 113

Although FPs are undoubtedly powerful tools for a variety of cell biological applications, as
researchers have pushed the boundaries to longer-term imaging and more specialized subcel-
lular compartments, limitations of the existing toolset have been uncovered. For example, long-
term expression of red FPs often leads to the accumulation of puncta that colocalize with
lysosomes, perhaps due to the resistance of highly stable RFPs to lysosomal proteases [20–23].
Many FPs characterized as monomeric have been shown to interact when expressed as a fusion
to membrane proteins [24,25]. Finally, inadvertent glycosylation sites and cysteine residues can
lead to the formation of intermolecular disulfide bonds, leading to higher-order oligomers and
interfering with chromophore formation, potentially causing mislocalization, particularly in the
crowded and oxidizing environment of the secretory pathway [23]. Some recent engineering
efforts have attempted to alleviate these limitations. For example, FusionRed was designed from
mKate with mutations that reduce toxicity, improve performance in fusions, and enhance
monomericity [22], and cysteine-less blue, cyan, green, and yellow FPs (oxFPs) have been
developed for use in oxidizing compartments [23].
Infrared FPs and Indicators: Advantages and Caveats
Long-wavelength light between 650 and 900 nm penetrates the furthest through animal tissue
because the combined effects of tissue absorbance (i.e., from hemoglobin, water, and lipids)
and light scattering are at a minimum [26,27]. Accordingly, infrared FPs (IFPs) are preferred for
use as protein tags and genetically encoded indicators for in vivo imaging applications [13].
Given that avGFP homologs with excitation in the near-infrared range have been neither
discovered in nature nor engineered in the lab, researchers have turned to bacterial phyto-
chromes (BphPs) as templates for engineering IFPs (Figure 1C). BphPs belong to the phyto-
chrome red/far-red photoreceptor superfamily [28,29] and typically exhibit maximum
absorbance at approximately 650–700 nm, but are natively not fluorescent. In nature, the
chromophore of BphPs is a covalently bound biliverdin (BV), a linear tetrapyrrole that is produced
by heme oxygenase 1 (HO-1) as the chief catabolic metabolite of heme. This strategy builds
upon the pioneering work of Lagarias and coworkers, who developed red and near-infrared
fluorescent ‘Phytofluors’from phytochrome proteins [12,30]. In contrast to BV-binding BphPs,
the earlier Phytofluor proteins required linear tetrapyrrole molecules, such as phycoerythrobilin
and phycocyanobilin, which are not produced by mammalian cells.
The first example of utilizing BphPs in mammalian imaging was IFP1.4, which was engineered
from a truncated DrBphP from Deinococcus radiodurans [13]. The utility of IFP1.4 was demon-
strated by fluorescence imaging of the liver in intact mice. Since thousands of bacteriophyto-
chrome-like sequences have been reported, this work opened a new door in engineering long-
wavelength FPs. Indeed, soon after, iRFP was engineered from RpBphP2 from Rhodopseu-
domonas palustris and used for imaging of mouse liver with improved brightness [31]. iRFP has
similar molecular brightness to that of IFP1.4, but is significantly brighter in cells and, thus,
appears to utilize endogenous BV better than does IFP1.4. IFP2.0, the improved version of
IFP1.4, was demonstrated to have similar brightness to that of iRFP when imaging brain tumors
in intact mice [32]. Yet other IFPs include: Wi-Phy, derived from truncated DrBphP [33]; iRFP720,
a red-shifted iRFP mutant; iRFP670 and iRFP702, both derived from RpBphP6 [34]; mIFP,
engineered from BrBphP (see below) [35]; and iBlueberry, a rationally designed mIFP variant with
a40-nm blue shift in both excitation and emission [36].
To optimally serve as a protein fusion tag, an IFP should be monomeric so as not to perturb the
stoichiometry of the protein of interest, which may affect protein function and/or trafficking.
Based on sequence and structural analysis of the dimer interface, a bacteriophytochrome from
Bradyrhizobium that is monomeric in its truncated form was identified from a sequence data-
base, and was engineered into a naturally monomeric IFP (mIFP) [35]. mIFP can be used to label
various proteins in live cells, Drosophila, and zebrafish, and, unlike dimeric iRFP, can be used to
114 Trends in Biochemical Sciences, February 2017, Vol. 42, No. 2

efficiently label the fine structure of dendrites in Drosophila. However, while mIFP is useful for
tagging and imaging proteins, it is about five times less photostable compared with iRFP.
Generally speaking, this is not a limiting factor for whole-animal imaging since the low-excitation
light intensities typically used [36] permit the acquisition of tens of thousands of images within the
photobleaching half-time of mIFP. However, the relatively low photostability can limit the duration
of time-lapse imaging experiments with mIFP-tagged proteins in live cells. Accordingly, in
addition to improving the intrinsic brightness, a future direction for mIFP development should
focus on increasing the photostability of monomeric IFPs or engineering a monomeric iRFP for
protein labeling.
One caveat of BphP-derived IFPs is that the fluorescence depends on BV concentration, which
varies in different cells and organisms. In Drosophila and zebrafish, BV concentration is limited
[32,35]. In mammals, neurons appear to have low concentration of BV. Thus, usage of BphP-
derived IFPs is limited in these organisms and cell types. However, this may provide an
opportunity for inducible fluorescence by delivering exogenous BV, which might open new
applications, such as pulse-chase labeling. Other than the exogenous addition of BV, one way of
overcoming this limitation is co-expression of HO-1, which converts heme into BV. For example,
HO-1 co-expression improves IFP2.0 fluorescence in Drosophila, mice brain tumors [32], and
zebrafish [35]. HO-1 has been reported to have anti-inflammatory and antioxidative functions via
its products, and its expression is highly induced in response to pathophysiological stress [37].
Although HO-1 overexpression may introduce biological perturbation, such toxicity has not been
observed in Drosophila and zebrafish. Nevertheless, an appropriate control is recommended
when BphP-derived IFPs are used in cells and animals even without overexpression of HO-1.
This is because expression of BV-binding IFPs may perturb the endogenous pool of BV and BV-
derived bilirubin, which is lipophilic and has antioxidant and cytoprotective roles in protecting
lipids from oxidation, complementing the water-soluble glutathione that protects water-soluble
proteins [38].
Since BphP-derived IFPs have a completely different protein structure (Figure 1C) to that of the
11-stranded b-barrel coelenterate FPs (Figure 1A,B), IFPs provide new opportunities for engi-
neering genetically encoded indicators. An infrared fluorogenic executioner caspase reporter
(iCasper) was recently developed, based on the unique interactions between mIFP and its
chromophore [39] (Figure 3). iCasper was used in imaging embryonic development in Drosophila
and revealed interesting spatiotemporal coordination between cell apoptosis and embryonic
morphogenesis, as well as dynamics of apoptosis during tumorigenesis in the brain of Dro-
sophila [39]. The iCasper technology should be easily adaptable for various protease activities,
CC
G
N
PAS
GAF
DEVDG
N00:00 00:18 00:36 00:42
00:54 01:00 01:02 01:10
PAS
GAF
DEVD
Caspase-3
(A) (B)
Figure 3. Design of an Infrared Fluorescent Protein (IFP) Protease Indicator. (A) Cartoon showing the cleavage and activation of iCasper. iCasper enables
imaging of caspase-3 activation during apoptosis, one type of programed cell death. (B) iCasper enables the visualization of the spatiotemporal dynamics of apoptosis in
vivo. Time-lapse images of neurons in the ventral nerve cord in Drosop hila reveal caspase activation and apoptotic cell shape change during central nervous system (CNS)
development. The neuron undergoing apoptosis is indicated with the arrow. The number at the top-right corner of each panel refers to time (h:min). Note that the neurons
also express membrane-targeted green fluorescent protein (GFP), which is brighter than the split GFP fluorescence from iCasper. Scale bar = 10 mm.
Trends in Biochemical Sciences, February 2017, Vol. 42, No. 2 115

enabling the dissection of signaling pathways that regulate protease activity, high-throughput
screening of protease inhibitors for drug development, and in vivo biological studies. The
development of the iCasper technology demonstrates that BphP-derived IFPs provide a
new and promising scaffold for designing fluorogenic indicators (e.g., of kinase activity and
membrane potential) that will be ideal for visualizing the spatiotemporal dynamics of cell signaling
in vivo.
A New Class of Far-Red FPs based on the Allophycocyanin a-Subunit from
A Cyanobacterial Phycobiliproteins
In an effort to further expand the color palette of FPs and improve upon the relativelylow quantum
yields and poor stability [13,31] of BphP-derived IFPs, a new class of FP was recently developed
from an allophycocyanin /-subunit (APC/) from the cyanobacterium Trichodesmium erythraeum
[40]. Native APC is a highly fluorescent hexamer that requires an auxiliary lyase to incorporate
phycocyanobilin (PCB). To develop a useful FP from native APC, the protein was engineered to be
self-sufficient (i.e., no lyase required) to covalently incorporate a BV rather than a PCB chromo-
phore. The protein was then subjected to 12 rounds of directed evolution for bright fluorescence
and low cytotoxicity to Escherichia coli. This effort ultimately led a bright APC/-derived FP that
was designated ‘small Ultra-Red FP’(smURFP) [40] because its light-blue color when viewed
under white light is reminiscent of the ‘Smurf’cartoon characters (Figure 4A). [3_TD$DIFF]smURFP is a
homodimer of 15-kDa subunits (Figure 1D) and has excitation and emission maxima at 642 nm
and 670 nm, respectively. [3_TD$DIFF]smURFP has an exceptionally large extinction coefficient
(e= 180 000 M
–1
[9_TD$DIFF]cm
–1
) and a modest quantum yield (F= 0.18), resulting in an intrinsic fluores-
cent brightness that is similar to that of EGFP. Accordingly, smURFP is the brightest far-red
(conventionally defined as FPs with emission maxima between 633 and 670 nm) or near-infrared
(emission maximum greater than 670 nm) FP yet reported (Figure 2B). Due to the limiting
concentration of BV in mammalian cells [41], treatment of transfected cells with 1–5mMBV
dimethyl ester (BVMe
2
) is typically necessary to obtain the brightest fluorescence. Yet other
favorable properties of smURFP include extremely good photostability and good expression in
neurons (Figure 4B) with no apparent formation of fluorescent puncta.
Using smURFP and IFP2.0, a far-red/near-infrared fluorescent ubiquitination-based cell cycle
indicator (FUCCI) [42] was created (Figure 4C). This new indicator could be combined with the
previously reported visible FP-based FUCCI for imaging cell cycle progression in two distinct cell
types [40].[3_TD$DIFF]smURFP expressed in HT1080 tumor rodent xenografts showed fluorescence even
(A) (B) (C) (D)
Figure 4. Small Ultra-Red FP ([3_TD$DIFF]smURFP): A Far-Red, Biliverdin-Binding Fluorescent Protein Derived from the Allophycocyanin /-Subunit from the
Cyanobacterium Trichodesmium erythraeum.(A) Purified smURFP (left) shows color reminiscent of the cartoon character's (Smurf) skin. (B) [3_TD$DIFF]smURFP expressed in
primary rat neurons. (C) Far-red/near-infrared fluorescent ubiquitination-based cell cycle indicator (FUCCI) expressed in HEK293A cells. Red-tinted fluorescence
corresponds to smURFP, which illuminates the G
0
/G
1
phases, while green-tinted fluorescence corresponds to IFP2.0, which illuminates the S/G
2
/M phases. (D) HT1080
tumor xenografts stably expressing smURFP. Fluorescence is visible without injection of exogenous biliverdin (BV).
116 Trends in Biochemical Sciences, February 2017, Vol. 42, No. 2

without the addition of exogenous BV (Figure 4D), although the fluorescence intensity was less
than with mCherry or mCardinal under similar conditions.
A Bilirubin-Inducible FP from the Vertebrate Subphylum
Recent years have seen the increasing recognition of the prevalence of biological fluorescence in
vertebrates. One such vertebrate is the Japanese eel, Anguilla japonica [43], which exhibits
green fluorescence from its skeletal muscle. The gene responsible for this green fluorescence
was cloned [14] and found to encode a 16.9-kD polypeptide in the fatty-acid binding protein
(FABP) family [44–46]. The protein, which was named UnaG (Figure 1E), had no intrinsic
fluorescence but showed bright green emission in mammalian cells and biological mixtures
even under anaerobic conditions. Extensive screening revealed the chromophore to be bilirubin,
a lipophilic bilin that is the reduction product of BV, a clinical diagnostic for liver function, and
responsible for the diseases jaundice and kernicterus [47–49]. UnaG binds to bilirubin non-
covalently with high affinity (K
d
= 98 pM) and high specificity. These favorable properties enabled
UnaG to be used to quantify serum levels of bilirubin from humans [14].
As an alternative to avGFP for live cell fluorescence imaging, UnaG has the inherent advantages
of a smaller gene size (which may facilitate packaging in viral vectors with limited DNA capacity),
and the ability to develop fluorescence in anaerobic conditions when bilirubin is supplied from
serum. Accordingly, UnaG fills an important niche in the toolbox of genetically encoded
fluorophores and is likely to serve as the prototype for a growing class of fluorescence-based
indicators [50–52].
Harnessing FP Photochromism for Super-resolution Microscopy or
Enhancing Contrast
While most FP-engineering efforts have focused on making improved tools for traditional
fluorescence microscopy, there has been a parallel effort to develop FPs that switch between
molecular states (photochromism). FP photochromism was first demonstrated in 1997 by
Moerner, Tsien, and colleagues, who established that single molecules of yellow FPs exhibited
intermittent fluorescence emission (i.e., blinking) and that molecules trapped in a long-lived dark
‘off’state could be converted back into the ‘on’state by illumination with high-energy (405 nm)
light [53]. Subsequent spectroscopic studies have largely reinforced the model of dark-state
conversion proposed by Moerner and colleagues, and have revealed multiple mechanisms for
excited-state and dark-state transitions, including conformational dynamics of the chromo-
phore, such as cis-trans isomerization [54,55], excited-state proton transfer, excited-state
solvation dynamics [56–58], and triplet state conversion [59]. However, it was not until 2002
that mutagenesis was first used to accentuate this photochromism, leading to the first designed
photoactivatable avGFP (PA-GFP) [60]. The notion that proteins could be engineered for
improved photochromism helped pave the way for the rapid expansion of super-resolution
microscopy platforms that exploit transitions between molecular states to localize molecules
with high precision.
Since the original development of microscopies that rely on the switching of molecules between
different molecular states to improve spatial resolution, such as stimulated emission deple-
tion (STED) [61,62],photo-activated localization microscopy (PALM) [63], and stochastic
optical reconstruction microscopy (STORM) [64], there has been a rapid proliferation of
techniques that exploit the complex photochromism of FPs to improve spatial resolution and/or
enhance fluorescence contrast (Table 1). Photochromism in FPs can be reversible or irreversible,
inducible or spontaneous, and can occur on a range of timescales, from 1-ms recovery from
triplet states [59] to 10–1000-ms recovery from transient dark states that can be either photo-
protective or photoreactive [65], or even much slower recovery from kinetically trapped dark
states from which irradiation can induce transition back to bright state [53]. Not surprisingly,
Trends in Biochemical Sciences, February 2017, Vol. 42, No. 2 117

different modalities exploit different photophysical properties, and have different requirements
for optimal performance, as outlined in Table 1. While several FPs have been explicitly engi-
neered based on irreversible photochromism for PALM/STORM requirements [66–69], only a
few FPs have been screened for microscopies that exploit reversible photochromism [70,71].
Sophisticated screens could be developed to explore the parameter space [72,73], perhaps
leading to greater insight into the mechanism(s) of these processes and probes optimized for
specific imaging modalities.
Resolving Biochemical Activities in Super-resolution
It has become increasingly clear that biochemical activities within the cell are often spatially
compartmentalized into regions, with sizes as small as tens of nanometers, known as micro- or
nanodomains [74]. While a large number of FP-based indicators have been developed to track
these biochemical events in living cells using standard fluorescence microscopy [75,76], there
are now a growing number of examples in which fluorescent indicators have been paired with
super-resolution imaging methods to produce cellular maps of biochemical activities in high
resolution.
A typical fluorescent indicator comprises a ‘sensing element’to detect the target biochemical
activity and a ‘reporting element’to translate the biochemical event into a change in the
fluorescent readout [77]. Depending on the reporting element, the indicators can be categorized
into different classes, such as those based on the translocation of fluorescence (Figure 5A),
change in Förster resonance energy transfer (FRET) efficiency (Figure 5B), and change in the
fluorescence of a single fluorescent protein (Figure 6A).
Our understanding of phosphoinositide (PI) dynamics has benefited from a series of indicators
designed to track the changes in various PI species in individual cells [78]. These probes, which
generally comprise a standard FP (the reporting element) fused to a protein domain that
specifically binds a PI (the sensing element), typically rely upon translocation of the probe to
the specific membrane compartment to report on the generation of the PI species under study.
Given the controversial model of lipid rafts and interest in probing lipid nanodomains in the
plasma membrane [79], it is tempting to use these PI indicators to map PI distribution in the live
Table 1. Fluorescence Imaging Modalities that Exploit FP Photochromism
Microscopy Photochromism Commonly
used FPs
Desired [5_TD$DIFF]properties for [6_TD$DIFF]optimal
[7_TD$DIFF]performance
Refs
PALM/STORM Photoactivation,
photoconversion,
reversible
photoswitching
mEos3.2,
mMaple
2or3,
PA-mCherry
High number of photons emitted/
switching cycle; ratio of rate constant
for on/off switching
[66–68]
RESOLFT Reversible
photoswitching
rsEGFP2 Rapid switching kinetics between
on/off states; high contrast ratio
between on/off states; number of
switching cycles before photobleaching
[70]
Stochastic optical
fluctuation imaging
(SOFI), pcSOFI
Spontaneous or
light-induced
dark-state
conversion
Dronpa,
rsTagRFP,
Skylan-S
Fluorescence fluctuations that are slow
compared with timescale of acquisition
time (tens of ms); photostable enough
to allow visualization of fluctuations over
duration of experiment
[71,163–165]
SAFIRe Dark-state
lifetime
Long-lived dark state with red-shifted
absorption
[166,167]
OLID, OPIOM Photoswitching
kinetics
Photoswitching dynamics matched
to modulation of excitation light
[168,169]
118 Trends in Biochemical Sciences, February 2017, Vol. 42, No. 2

cell membrane under high resolution, but this task is not trivial. One of the complicating factors is
that direct observation of lipid clusters at a nanometer-length scale is hampered by the diffraction
of light. To overcome this limitation, researchers have combined FP-based PI indicators and
super-resolution imaging to quantify PIs in the plasma membrane and visualize their distribution.
In a recent study [80], PALM was used to visualize and quantify phosphatidylinositol 3-phos-
phate (PI3P), a lipid involved in endocytosis and membrane transport. A photoswitchable FP,
mEos2 [81], was fused to a PI3P-binding domain (a double FYVE domain of EEA1) to construct a
super-resolution compatible PI3P indicator. Although colocalization experiments using different
markers suggested that mEos2 did not alter the localization pattern of the FYVE domain in this
case despite its propensity to dimerize [68], the nondimerizing variant mEos3.2 should provide a
better alternative to eliminate any potential dimerization artifacts. In a separate study, van den
Bogaart et al. used a phosphatidylinositol 4,5-bisphosphate (PIP
2
) indicator (the pleckstrin
homology domain of protein lipase C delta fused to [18_TD$DIFF]Citrine) in combination with STED micros-
copy, to map PIP
2
clusters on the plasma membrane of a cell [82].
Perhaps the most widely used class of indicators is those developed for tracking protein–
protein interactions. While many of these indicators are based on FRET, protein fragment
complementation [83] is another popular approach. Protein fragment complementation
requires two nonfluorescent FP fragments that do not self-associate on their own, but that
can recombine to form a functional FP if brought into close proximity with the help of an
interacting pair of proteins. This property, which is the basis for bimolecular fluorescence
(A)
(B)
Cytosolic
fluorescence
Plasma
membrane-localized
fluorescence
Low FRET High FRET
Kinase
CFP
Phosphatase P
Substrate
PAABD
YFP
Figure 5. Schematic Representations of Indicators for Imaging of Intracellular Signaling Activity. (A) Transloca-
tion-based indicator design. (B) Intramolecular Förster resonance energy transfer (FRET)-based design of a kinase activity
indicator.
Trends in Biochemical Sciences, February 2017, Vol. 42, No. 2 119

complementation (BiFC) and several related protein fragment complementationassays (PCAs),
has been exploited to detect protein–protein interactions in living cells and even whole animals
[84], although the effectively irreversible nature of BiFC has limited its application in tracking
dynamic interactions. To generate high-resolution maps of specificprotein–protein interac-
tions, specific pairs of fragments have been identified for photoactivatable or photoswitchable
proteins and used for imaging protein–protein interactions in super-resolution [85–88].For
example, a BiFC-PALM approach was recently developed using PAmCherry1 [87].Nickerson
et al. studied interactions between the small GTPase Ras and its downstream effector Raf using
this approach and observed nanoscale clustering and diffusion of individual KRas G12D/CRaf
RBD (Ras-binding domain) complexes on the cell membrane [87]. The Zhang laboratory
recently described a strategy in which protein–protein interactions induce the reconstitution
of fluorescent proteins capable of exhibiting single-molecule fluctuations suitable for stochastic
optical fluctuation imaging (SOFI). Subsequently, spatial maps of these interactions can be
resolved in super-resolution in living cells based on statistical analysis of the fluorescence
fluctuations. This strategy, termed ‘reconstituted fluorescence-based SOFI’(refSOFI), was
used to investigate the interaction between the endoplasmic reticulum (ER) Ca
2+
[17_TD$DIFF] sensor STIM1
and the pore-forming channel subunit ORAI1 at the ER–plasma membrane junction [86].
Limitations of these BiFC-based approaches include irreversibility and long maturation time,
ranging from 30 min to hours. It can be anticipated that new indicators with better temporal
resolution and reversibility will continue to emerge to help reveal a more detailed map of
biochemical activities in cells.
Low fluorescence High fluorescence
Ca2+/CaM-
binding
peptide
Ca2+/CaM-
binding
peptide
Ca2+
cp linker cp linker
CaM
Ca2+/CaM
Ca2+/CaM
Ca2+/CaM
GFP RFP
Chromophore Chromophore Lys78Arg376 Cterm
Nterm
Cterm
Nterm
(A)
(B) (C)
Figure 6. Single Fluorescent Protein (FP)-Based Genetically Encoded Calcium Ion (Ca
2+
) Indicators (GECIs).
(A) Schematic representation of mechanism of response of a GCaMP-type Ca
2+
indicator. (B) Structure of GCaMP6m
[Protein Data Bank (PDB) ID 3WLD) [170]. Arg376 modulates the fluorescent response upon binding to Ca
2+
by electrostatic
stabilization of the anionic state of the chromophore. (C) Structure of R-GECO1 (PDB ID 4I2Y) [105]. Lys78 is proposed to
act similarly to Arg376 of GCaMP6m for modulation of the Ca
2+
-dependent fluorescent response.
120 Trends in Biochemical Sciences, February 2017, Vol. 42, No. 2

Genetically Encoded Calcium Ion Indicators
Paralleling the development of improved FPs and indicators of biochemical activities have been
concerted efforts to develop improved genetically encoded calcium ion (Ca
2+
) indicators
(GECIs). The first examples of single FP-based Ca
2+
indicators (Figure 6A) were reported in
1999 [89], just 2 years after the debut of the FRET-based ‘cameleon’-type indicators [90].
Cameleon-type indicators produce a ratiometric fluorescent signal as a result of Ca
2
+
-dependent changes in the efficiency of FRET from a blue-shifted donor to a red-shifted
acceptor. By contrast, single FP-based indicators produce Ca
2+
-dependent fluorescent
changes (typically, but not always, an intensiometric fluorescent signal) as a result of
modulation of the proteinaceous environment of the chromophore. The first-generation single
FP-based indicators (‘camgaroo’type) were created through the clever insertion of calmodulin at
position 145 of EYFP [89]. In camgaroo, the conformational change associated with the binding
of Ca
2+
to calmodulin modulated the chromophore environment such that the pK
a
shifted to a
lower value and the fluorescence intensity increased sevenfold. The second generation of single
FP-based Ca
2+
indicators, specifically the Pericam construct from Miyawaki [91] and the G-
CaMP construct from Nakai [92], exploited a circularly permuted FP topology that had been first
predicted by Tsien and coworkers [89]. These second-generation indicators comprised an FP
that was circularly permuted such that the new termini were in close proximity to the chromo-
phore. Fused to the termini were calmodulin and a calmodulin-binding peptide that underwent a
Ca
2+
-dependent interaction and caused the requisite modulation of the chromophore
environment.
Early efforts to further improve single FP-based Ca
2+
indicators, specifically the G-CaMP design
[92], were hampered by the lack of a protein structural model, or even a solid hypothesis about
the mechanism by which the indicator functioned [93]. Two independent reports of the X-ray
crystal structure of a second-generation variant [94,95] served as a catalyst for future structure-
guided engineering efforts, soon facilitating the development of GCaMP3 [96]. GCaMP3 is
generally considered to be the ‘breakthrough’version that was practically useful for routine
neuronal activity imaging in a variety of contexts, including transgenic mice [97]. A series of
additional improved versions have since been reported [98–101]. The current state-of-the-art
variants are the GCaMP6 (Figure 6B) series produced by the Genetically-Encoded Neuronal
Indicator and Effector (GENIE) project at the Howard Hughes Medical Institute (HHMI) Janelia
Research Campus [102]. Transgenic mice expressing fast and slow variants of GCaMP6 under a
neuron-specific Thy-1 promoter are now available [103].
In addition to serving as the catalyst for rapid improvements in the GCaMP series, the X-ray
crystal structures of GCaMP2 [94,95] also helped accelerate the development of variants with
altered fluorescent hues. In 2011, the Campbell lab reported a series of new indicators, including
blue, blue-green emission ratiometric, and red fluorescent variants, which they designated the
GECO-series [104]. Parallel efforts from the GENIE project also led to a series of color variants
that included blue, cyan, yellow, and red Ca
2+
indicators [105]. The most promising new
indicators to arise from these efforts were the two red fluorescent variants: mApple-derived
R-GECO1 from the Campbell lab (Figure 6C), and mRuby-derived RCaMP from the GENIE
project. Generally speaking, red fluorescent indicators are preferable to green ones due to their
lower autofluorescence, lower phototoxicity, and greater tissue penetration associated with
longer wavelength excitation. As with the GCaMP series, the red indicators have continued to be
improved [106,107], with the latest variants (i.e., the R-GECO1-derived R-CaMP2 [108], and
jRCaMP1a,b and jRGECO1a variants from the GENIE project [109]) offering performance that
approaches that of GCaMP6.
The toolbox of single FP-based indicators continues to expand in several directions, with other
engineering efforts focused on making improved yellow indicators [110], a long Stokes-shift
Trends in Biochemical Sciences, February 2017, Vol. 42, No. 2 121

indicator [111], variants with lower K
d
values that are suited for imaging Ca
2+
in the ER [112,113],
and photoconvertible variants that can be selectively ‘highlighted’by spatially defined conversion
to a spectrally distinct fluorescence hue [114,115]. Interestingly, it was recently reported that
GCaMP6 itself can be ‘highlighted’by photoconversion to a red state that retains Ca
2+
responsiveness [116]. An exciting twist on the utility of photoconvertible Ca
2+
indicators is
the recently reported CaMPARI variant [117]. CaMPARI is a GCaMP-type indicator that exhibits
Ca
2+
-dependent photoconversion to a red state and, thus, acts as an integrator of neuronal
activity during a period of illumination.
FP-Based Indicators of Transmembrane Voltage
While GECIs are the current workhorse of neuronal activity imaging, neuroscientists have long
recognized the need for effective genetically encoded voltage indicators (GEVIs). Neurons
process and transmit signals (an action potential) via changes in their transmembrane voltage.
A negative-inside transmembrane voltage is established across the membrane by ion pumps
and transporters, and can be modulated with millisecond kinetics by the action of neurotrans-
mitter- and voltage-gated ion channels [118]. If voltage changes could be visualized in real time
with a fluorescent indicator, then neuroscientists would be able to study questions such as how
inputs at different synapses are integrated to induce action potential firing, and how neurons in a
circuit fire in a coordinated manner. One major challenge for optical imaging of voltage changes
is that a single action potential is complete within a few milliseconds and so image acquisition
rates in the range of 100 to 1000 Hz are essential [118]. The correspondingly short image
acquisition times and high-intensity excitation light sources necessitate that practically useful
voltage indicators be particularly bright and photostable. Further complicating the use of voltage
indicators is the fact that an indicator of membrane potential must be confined to the 2D surface
of the plasma membrane, rather than the 3D volume of the cytoplasm. By contrast, elevated
cytoplasmic Ca
2+
levels persist for hundreds of milliseconds following an action potential [119],
which enables relatively facile detection of these changes with much higher signal:noise ratios
than are typically achievable for voltage indicators.
The field of GEVI development is as old as the field of GECI development, but has matured at a
slower rate due to the more challenging nature of GEVI design (Figure 7) and optimization. The
first GEVI, FlaSh, comprising a GFP domain inserted in an intracellular segment of a voltage-
gated potassium channel, was published in 1997 [120], the same year as the first GECI,
Cameleon [90]. FlaSh exhibits a 5% decrease in GFP fluorescence upon membrane depolari-
zation in the physiological range of –70 to +30 mV, but with rather slow kinetics. Subsequently,
SPARC, a GFP insertion in an intracellular loop of a voltage-gated sodium channel, was found to
show small (<0.5%) changes in fluorescence upon depolarization but with fast kinetics (0.8-ms
activation time-constant) [121]. In both FlaSh and SPARC, the mechanism of voltage sensing by
the GFP is unknown. In addition, a GEVI named VSFP1 was created by fusion of CFP and YFP in
tandem at the C terminus of an isolated voltage-sensing domain (VSD) from a voltage-gated
potassium channel [122]. VSFP1 shows a 2% increase in YFP fluorescence upon depolarization
with fast kinetics (0.7-ms activation time-constant), presumably because changes in the distri-
bution of FP orientations upon voltage-induced VSD movements induce an increase in FRET.
However, poor membrane expression of FlaSh, a FlaSh derivative named Flare, and VSFP1 in
mammalian neurons prevented their actual use in neuroscience applications [123].
The discovery of voltage-sensing phosphatases in 2005 [124] provided a VSD that was well
expressed at the membrane and also generated larger changes in the fluorescence of attached
FP domains, serving as the basis for a series of increasingly effective voltage indicators of
different architectures [125]. Fusion of this new VSD to CFP and YFP at the C terminus (VSFP2),
or to one FP at the N terminus and another at the C terminus (VSFP-Butterfly), gave 10%
fluorescence changes with fast (1–3 ms) kinetics [126,127]. In addition, fusions of single FP
122 Trends in Biochemical Sciences, February 2017, Vol. 42, No. 2

domains at the C terminus (VSFP3s) showed small (1.6–3.5%) and fast (1.8–3.8 ms) fluores-
cence decreases upon depolarization [128]. ArcLight, a fusion of VSD and a pH-sensitive FP with
an Ala-to-Asp mutation on the b-barrel surface, shows a larger (35%) but slower (10 ms)
fluorescence decrease [129]. ASAP1, an insertion of a circularly permuted GFP in an extracellular
loop of the VSD, shows both large (25%) and fast (2 ms) fluorescence decreases upon
depolarization [130]. Finally, Abdelfattah et al. recently reported FlicR1, a bright and fast red
fluorescent voltage indicator based on fusion of a circularly permuted RFP to the C terminus of a
VSD, that exhibits fluorescence increases of 3% for a single action potential [131].
Yet another GEVI design is based on the genetic fusion of an FP to an opsin protein. Here, the FP
serves as a FRET donor to the opsin [132]. Depolarization increases opsin absorption at orange
wavelengths, increasing FRET from the FP, and decreasing FP brightness [133,134]. Among
these ‘FRET-opsin’GEVIs, the brightest and fastest are the Ace-mNeon series [135].
(A) FlaSh/Flare/SPARC
VSFP-Buerfly
ASAP1
VSFP3/ArcLight
VSFP1/VSFP2
(C)
(E)
(B)
(D)
out (+)
(+)in (–)
out (+)
in (–)
out (+)
in (–)
out (+)
in (–)
out (+)
in (–)
(–)
(+)
(–)
(+)
(–)
(+)
(–)
(+)
(–)
GFP GFP
CFP YFP CFP YFP
FRET
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
YFP RFP YFP RFP XFP
XFP
cpGFP
cpGFP
FRET
Figure 7. Designs of Fluorescent Protein (FP)-Based Genetically Encoded Voltage Indicators (GEVIs). (A) In FlaSh/Flare/SPARC, green fluorescent protein
(GFP) is inserted in an intracellular segment of a homotetrameric potassium channel (FlaSh/Flare) or of a pseudotetrameric sodium channel (SPARC). Only one repeat of
the six-helix transmembrane motif is shown for clarity. (B) In VSFP1/2, a FRET pair is fused to the intracellular segment following a four-helix voltage-sensing domain
(VSD). (C) VSFP-Butterfly comprises a FRET pair of FPs (YFP and RFP) fused to the termini of the VSD. (D) In VSFP3/ArcLight, a single FP is fused following the VSD. (E) In
ASAP1, a circularly permuted GFP (cpGFP) is inserted into an extracellular loop of the VSD.
Trends in Biochemical Sciences, February 2017, Vol. 42, No. 2 123

The utilization of FP-based GEVIs is still in its infancy. VSFP-Butterfly was used to visualize rapid
synchronized voltage responses in neuronal populations in the mouse sensory cortex [127].
ArcLight detected odorant-induced depolarizations and hyperpolarizing responses (which are
undetectable with calcium sensors) in living flies [136]. ASAP2f was used to deduce differences
in voltage-calcium coupling in the fly visual system [137]. Ace-mNeon has been used to visualize
subcellular voltage changes in mouse cortex and to measure voltage propagation rates in fly
olfactory neurons [135]. A remaining challenge is to use FP-based GEVIs for temporally precise
monitoring of voltage transients in individual neurons in the mammalian brain, where two-photon
excitation would be helpful for background suppression and fast-scanning approaches will be
required for temporal resolution.
Optogenetics with FPs
An optogenetic actuator is a genetically encoded protein that undergoes an illumination-
dependent change in function that induces, disrupts, or otherwise changes, a cellular function.
While channelrhodopsin-2, the blue-light activated cation channel, is often considered the
archetypical optogenetic actuator [138], FPs were in fact some of the earliest optogenetic
actuators developed. The earliest examples of FP-based optogenetics were based on the
principles of chromophore (which could also be a fluorophore)-assisted light inactivation (CALI)
(Figure 8A). The mechanism of CALI involves the photochemical generation of reactive oxygen
species (ROS) by the triplet state of the excited chromophore [139,140]. ROS generated during
illumination can diffuse over short distances and oxidize specific amino acid residues of nearby
proteins, leading to crosslinking and photo-induced cleavage [141]. Due to the reactive nature
and short lifetime of the ROS, CALI is typically specific to the protein or protein complex to which
the chromophore is tethered. This photodestructive approach can be used to irreversibly knock
out protein functions and disrupt cellular pathways. This CALI effect was observed with EGFP
fusion proteins and was used to inhibit galactosidase [142], focal adhesion kinase [143], MyoII
regulatory light chain [144], and Connexin 43 [145]. EGFP is a relatively poor mediator of CALI
Enzyme/
protein of
interest
Enzyme/
protein of
interest
Enzyme/
protein of interest
Enzyme/
protein of interest
490-nm light
390-nm light
Enzyme/
protein of
interest
Photosensizer Photosensizer Photosensizer
Light ROS
ROS
(A)
(B)
ROS
ROS
Dronpa
Dronpa
Dronpa
Dronpa
Figure 8. Two Current Approaches to Control Protein and/or Cellular Functions with Fluorescent Protein (FPs). (A) FPs capable of generating reactive
oxygen species (ROS) during illumination (photosensitizer), such as EGFP, miniSOG, KillerRed, and SuperNova, can be used to disrupt the function of protein(s) of interest
tethered to the photosensitizer. This is the principle of chromophore-assisted light inactivation (CALI). (B) Light-controll ed, reversible oligomerization of Dronpa can
sterically block the catalytic or functional site on fusion protein. This oligomerization process is controlled with two different wavelengths of light.
124 Trends in Biochemical Sciences, February 2017, Vol. 42, No. 2

due to the protected environment of the chromophore embedded within the b-barrel, and the
fact that EGFP was evolved to have high fluorescence quantum yield and a limited triplet state
[142]. Efforts to increase the ROS generation by FP under illumination has produced KillerRed
[146,147], a dimeric red FP that is capable of generating superoxide radical anion during green
light illumination [148,149]. Given that the dimeric nature of KillerRed complicates its use in CALI
experiments as a fusion protein, Takemoto et al. developed a monomerized version of KillerRed,
named SuperNova, which can be used for CALI and is less likely to perturb the localization of a
fusion protein [150].
MiniSOG is an engineered variant of a FMN-binding light, oxygen, voltage (LOV) domain from
Arabidopsis thaliana that generates ROS during blue light illumination [151]. CALI with miniSOG
was used to inhibit synaptic proteins [152,153], disrupt the CaMKII analog UNC-43 in Caeno-
rhabditis elegans [154], and in experiments to ablate cells [155]. SOPP, a newer variant
engineered from miniSOG, is reported to have eightfold improved singlet oxygen quantum
yield, but is yet to be tested in living cells [156]. A newly discovered LOV domain from
Pseudomonas putida (Pp2FbFP L30 M) is reported to have singlet oxygen quantum yield that
is threefold higher than that of miniSOG [157], which may be a suitable template for future
development of flavin-binding FPs as optogenetic tools to alter cellular function.
Other than utilizing FPs for CALI and cell ablation experiments, the utilization of FPs to control the
activities of intracellular enzymes and proteins has been demonstrated [158]. For example, an
engineered variant of the photoswitchable Dronpa FP enables switching between a tetrameric
and monomeric state by illumination with different wavelengths of light (Figure 8B). By using light
to control the oligomerization state of this Dronpa mutant, it is possible to block the active site of
attached enzymes and proteases, leading to the optogenetic control of protein functions.
Currently, its use may be limited due to the concentration-dependent nature of the association;
however, the light-induced oligomerization and dissociation of Dronpa can be further exploited
to develop novel methods of optogenetic control of intracellular pathways.
With the rapid adoption of optogenetic approaches and recent advances towards achieving all
optical observation and manipulation of biological pathways [159], the potential utilization of FPs
for simultaneous observation and manipulation may further revolutionize the way intracellular
events are studied in the coming decades.
Concluding Remarks
The authors of this review are enthusiastic developers and users of FP technology, and one of us
(R.Y.T.) was among the first researchers to recognize the utility of engineered FPs as tools for
fluorescence imaging. During the mid-1990s, it was surprising and gratifying that color variants
of avGFP could be created. We have been astounded by the myriad applications that have been
enabled by FP technology over the past two decades. We have no reason to fear that FPs have
yet given up all their secrets, and it is a safe bet that the coming years will see further advances.
While some of the general directions for these advances are apparent (e.g., FPs from new
organisms, near-infrared fluorescence, new classes of indicator, and new interactions with
photons), history has taught us to expect the unexpected.
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