Newly engineered cyan fluorescent proteins with enhanced performances for live cell FRET imaging.

Biotechnology Journal

Biotechnol. J. 2014, 9, 180–191

DOI 10.1002/biot.201300198

www.biotechnology-journal.com

Review

Newly engineered cyan fluorescent proteins with enhanced performances for live cell FRET imaging Fabienne Mérola, Asma Fredj, Dahdjim-Benoît Betolngar, Cornelia Ziegler, Marie Erard and Hélène Pasquier CNRS, Université Paris Sud, Laboratoire de Chimie Physique, Orsay, France

Cyan fluorescent proteins (CFPs) derived from Aequorea victoria green fluorescent protein are the most widely used Förster resonant energy transfer (FRET) donors in genetically encoded biosensors for live-cell imaging and bioassays. However, the weak and complex fluorescence emission of cyan variants, such as enhanced cyan fluorescent protein (ECFP) or Cerulean, has long remained a major bottleneck in these FRET techniques. Recently, several CFPs with greatly improved performances, including mTurquoise, mTurquoise2, mCerulean3, and Aquamarine, have been engineered through a mixture of site-directed and large-scale random mutagenesis. This review summarizes the engineering and relative merits of these new cyan donors, which can readily replace popular CFPs in FRET imaging protocols, while reaching fluorescence quantum yields close to 90%, and unprecedented long, near-single fluorescence lifetimes of about 4 ns. These variants display an increased general photostability and much reduced environmental sensitivity, notably towards acid pH. These new, bright, and robust CFPs now open up exciting outlooks for fluorescence lifetime imaging microscopy and advanced quantitative FRET analyses in living cells. In addition, the stepwise engineering of Aquamarine shows that only two critical mutations in ECFP, and one in Cerulean, are required to achieve these performances, which brings new insights into the structural bases of their photophysical properties.

Received 30 JUN 2013 Revised 17 SEP 2013 Accepted 31 OCT 2013

Keywords: Aquamarine · Cerulean · FLIM (fluorescence lifetime imaging microscopy) · FRET (Förster resonant energy transfer) · mTurquoise

1 Introduction Genetic fusions with fluorescent protein variants (e.g. green fluorescent proteins (GFPs)) provide genetically encoded fluorescent reporters with high biocompatibility, specificity, and integration into cellular pathways [1, 2]. In particular, numerous Förster resonant energy transfer (FRET) biosensors have been developed to decipher the specific molecular activities involved in signal transduction, such as second messenger levels, enzymatic activi-

Correspondence: Dr. Fabienne Mérola, Laboratoire de Chimie Physique, Bat 349, Université Paris Sud, F-91405 Orsay, France E-mail: [email protected] Abbreviations: ECFP, enhanced cyan fluorescent protein; EYFP, enhanced yellow fluorescent protein, FLIM, fluorescence lifetime imaging microscopy; FRET, Förster resonant energy transfer; mTFP, monomeric teal fluorescent protein; RSFP, reversibly switchable fluorescent proteins

180

ties, or formation of protein complexes [3, 4]. These powerful bioanalytical techniques are now intensively used to understand diseases, screen drugs, or monitor therapeutic efficacy within live cells or whole organisms [4–7]. The FRET interaction provides a general means to engineer a specific optical response at the nanoscale into a wide variety of molecular devices. FRET occurs when a fluorescent donor molecule in its excited state transfers its excitation energy non-radiatively to a nearby acceptor [8, 9]. The FRET interaction requires an overlap between the donor emission and acceptor absorption spectra, close proximity, and proper orientation of the two molecules. The efficiency of the FRET interaction sharply depends on the relative spatial arrangement of the donor–acceptor pair, which underlies its exquisite sensitivity to conformational changes or molecular association. The FRET interaction manifests itself through various perturbations of the fluorescence signals, such as a decrease in the fluorescence intensity and lifetime of the

© 2013 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim


Biotechnol. J. 2014, 9, 180–191


donor, an increase in the sensitized acceptor fluorescence, or changes in its polarization state. Two categories of GFP-based FRET sensors must be distinguished. Single-chain FRET biosensors connect a donor and an acceptor GFP covalently around a core sensor module that is expected to undergo some conformational change or cleavage (intramolecular FRET). Intramolecular FRET is widely used to monitor the intracellular levels of various ions, metabolites, or enzymatic activities [3, 4]. Two-component FRET systems (intermolecular FRET) require double transfections with separate donor- and acceptorlabeled gene constructs. The FRET responses then monitors variations in the spatial proximity and interactions of the two proteins. Intermolecular FRET is mostly used to establish direct protein–protein connectivity and assembly in signaling cascades, but can also report on posttranslational modifications, such as protein phosphorylation [10], or changes in the activation status of small [11] or heterotrimeric [12] GTPase proteins

2 Quantification issues in FRET techniques As a chemical imaging technique, FRET involves advanced fluorescence detection protocols and imposes stringent requirements on the fluorescent probes used. In general, small intracellular FRET changes must be monitored as a function of time and space, and should specifically reflect the chemical activity under scrutiny; this requires a very good stability of the donor and acceptor fluorescence signals. While intermolecular FRET techniques must usually cope with great signal variability, due to the uncontrolled donor–acceptor expression levels in transiently transfected cells, single-chain FRET sensors have the advantage of fixing the donor–acceptor stoichiometry. On the other hand, any intrinsic environmental sensitivity of the fluorophores to temperature, pH, or any other cellular component, as well as short- or longterm photoreactions, are clearly undesirable. To evaluate these problems, negative controls with non-responsive forms of the FRET biosensor may sometimes be implemented, for example, by mutating a critical residue in the core sensing or interaction domains [13]. Most biological studies rely on dose–response analyses, which require the routine collection of quantified FRET data. Although many different types of FRET indices have been proposed, the FRET efficiency, defined as the fraction of donor fluorescence photons transferred to the acceptor, is the most useful and transferable parameter (discussed in [14]), and carries precise physical and chemical meanings. A very convenient FRET detection mode is ratiometry, which monitors the raw ratio of acceptor over donor fluorescence intensities, and can be implemented in real-time under wide-field video microscopy. Due to spectral bleed-through between donor and acceptor channels, the relationship of fluorescence inten-


www.biotecvisions.com

sity ratios to the FRET efficiency remains complex. Also, the method is not applicable in the case of large variations in the donor/acceptor expression levels. In techniques based on acceptor photobleaching, dequenching in the donor fluorescence is monitored upon local photodestruction of the acceptor. These methods can, in principle, provide absolute FRET efficiencies, but cannot be used conveniently for dynamic studies. An emerging technique called photochromic FRET, which is currently restricted to a few FRET pairs, takes advantage of the reversible photoswitching properties of some GFPs (see Section 5) to “turn off” the acceptor absorbance at will [15, 16]. Spectral FRET or three-cube FRET (where bleed-through corrections are used to recover the FRET efficiency), and fluorescence lifetime imaging microscopy (FLIM; using fluorescence lifetime variations to directly monitor the donor fluorescence quenching) are the most general methods for the dynamic monitoring of FRET efficiencies. All of these FRET techniques have been extensively discussed in the literature for their principles, relative merits, and pitfalls [9, 14, 17]. Regardless of the method of detection used, it is important to recall that samples containing both donor and acceptor fluorophores, either covalently connected or present as independent molecular entities, may also display, to some extent, what is frequently called “non-specific FRET”. Such a FRET signal arises primarily from the random proximity of donor and acceptor molecules at high concentration and becomes significant when the fluorophore concentration approaches the sub-millimolar range, which is a level that is easily attained by cytosolic proteins in transiently transfected cells [18], and moreover, when the proteins are targeted to specific subcellular compartments. This proximity FRET occurs regardless of the formation of any molecular complex and depends only on the acceptor concentration [18] or its surface density in membrane expression systems [19]. For a cytosolic expression of the enhanced cyan fluorescent protein (ECFP)/enhanced yellow fluorescent protein (EYFP) pair, a proximity FRET efficiency of about 2% is expected for 100 μM EYFP, and will follow an approximate linear relationship with the acceptor concentration [18]. This nonspecific contribution may be assessed by monitoring the FRET signal of appropriate control samples as a function of the acceptor concentration or surface density. Quantification remains a major challenge of modern biochemical imaging. Reliable FRET quantifications require fluorophores with highly specific and well-understood photophysical responses. This is crucial for the advanced determination of intracellular physicochemical parameters, such as absolute ion or metabolite concentrations, or the binding affinity and stoichiometry of functional protein assemblies.

181


Biotechnol. J. 2014, 9, 180–191 www.biotecvisions.com


3 Short history of CFP engineering: From early mutants of AvGFP to Aquamarine The numerous applications of FRET bioimaging have triggered intensive efforts in GFP development, leading to the discovery of many new colors and brighter fluorophores, and the proposal of several alternative FRET pairs operating in different spectral ranges [20–29]. Among those, the early cyan fluorescent protein (CFP)/ yellow fluorescent protein (YFP) combination proposed by Tsien [30] remains the most popular choice, with a lower estimate of more than 60% of current published FRET studies. Moreover, this donor–acceptor pair is still chosen in the vast majority of newly built FRET sensors and is also widely used for in vivo imaging purposes [31–34] or for establishing stable fluorescent cell lines [5]. Cyan and yellow reporters are essential for multicolor imaging and multiplexed sensing of complex signaling processes, such as to monitor third-party components or to allow the simultaneous use of several FRET biosensors [9, 35, 36]. Meanwhile, in-depth scrutiny of ECFP and EYFP variants has revealed their numerous drawbacks [20, 37–42]. Although several improved YFPs have been developed, with Citrine and Venus ranking among the brightest monomeric GFPs known today [1, 14, 26], the poor per-

formances of the available CFPs has remained a major concern. The ECFP variant (Table 1) is derived from the Y66W mutation (the so-called “W” mutants) introduced by Tsien and co-workers [43, 44] within the Ser65-Tyr66-Gly67 chromophore of AvGFP . This mutation resulted in a major increase in chromophore size, which required multiple rounds of further random mutagenesis and screening to restore substantial fluorescence [30]. The resulting, extensively mutagenized ECFP is probably the least optimal FRET donor that microscopists may have dreamed of, because it suffers from low brightness, limited photostability, and very complex photophysics with multiexponential fluorescence decays [45]. This complex photophysics not only precludes detailed quantitative exploitation of FLIM data [38], but also comes together with strong environmental responses, such as those to pH, temperature, or ATP [45, 46]. Improving the ECFP protein by genetic engineering has been the subject of longstanding efforts (Table 1). Early studies by NMR spectroscopy [47] and X-ray crystallography [48] suggested that the conformational flexibility of strand 144–149 in the CFP beta-barrel was a major cause of the complex ECFP fluorescence. These ideas directly inspired the sitedirected design of Cerulean [49], the photophysical

Table 1. Comparison of the photophysical properties of CFP variantsa)

CFP variant

Mutations (relative to ECFP)

ECFP

εMax (M–1cm–1) ± S.D. 3000

Φf

Fluorescence lifetimeb) (ns) ±S.D. 2%

Bleaching timec) (s) ± S.D. 15%

pK1/2d) ± 0.1

30 000

0.37

2.50

730

5.6

000e)

0.67

3.05

640

5.2

Cerulean

H148D S72A-Y145A

33

mTurquoise

H148D-T65S S72A-S175G A206K

33 000f)

0.84

4.06

1480

3.4

mCerulean3

H148G-T65S + 8 mutations

30 000g)

0.87g)

N.D.

N.D.

3.2g)

Cerulean-T65S

H148D-T65S S72A-Y145A

34 800

0.84

3.96

780

3.6

mTurquoise2

H148D-T65S S72A-S175G A206K-I146F

30 000h)

0.93h)

N.D.

N.D.

3.1h)

Aquamarine

H148G-T65S

26 000

0.89

4.12

900

3.3

a) All data from [51, 56], except otherwise indicated. N.D. = not determined under comparable conditions. b) Average lifetime, =Σciτi, obtained from high-resolution time-correlated single-photon counting (TCSPC) measurements of the purified proteins. c) Exponential time constants of the irreversible loss in fluorescence intensity under constant wide-field illumination at 0.2 W/cm2 on cytosolic proteins expressed in Madin Darby canine kidney (MDCK) cells (averages of N ≈ 20–30 decays) [51]. d) pH value at which the fluorescence intensity dropped by 50% of the intensity at neutral pH. e) From [49]. f) From [53]. g) From [54]. h) From [55].

182





improvements of which can be mostly traced to the mutation H148D [45, 50]. The increased fluorescence quantum yield and lifetime of Cerulean (Table 1) has motivated its adoption in many recent FRET protocols [6, 9, 33, 35]. However, Cerulean suffers from degraded photostability relative to ECFP [1], while it retains a complex photophysics with multiple fluorescence lifetimes [51]. Gadella and co-workers [50] intensively worked on the further improvement of CFPs and YFPs. Notably, they combined the H148D mutation in ECFP with a series of mutations borrowed from the yellow variant Venus [52], which was expected to improve protein folding and chromophore maturation. These efforts led to SCFP3A, which showed, besides improved bacterial expression, a slightly higher fluorescence (+15%) than that of Cerulean [50]. Four years later, the same group further engineered SCFP3A by site-directed saturation mutagenesis, leading to the selection of the T65S mutation [53]. This resulted in mTurquoise, displaying a fluorescence quantum yield of 84% (Table 1) and, for the first time for a tryptophan-based chromophore, a fluorescence emission closely approaching a single-exponential decay law. The development of mTurquoise was followed by a series of new CFPs with similar, near-ultimate performances (Table 1). First, Cerulean was extensively remodeled into mCerulean3, owing to large-scale, multisite random mutagenesis, with up to 10 final mutations away from ECFP [54]. We then showed that the single-point mutation T65S was sufficient to confer equivalent performances to Cerulean [51]. Meanwhile, mTurquoise was further improved to mTurquoise2 through structure-guided identification of the I146F mutation [55]. Finally, we showed that a glycine residue was the optimum choice at position 148 in ECFP which, in combination with the T65S mutation, led to minimally engineered, but highly optimized Aquamarine [56]. All of these improved CFPs not only open new perspectives to sensitive and quantitative FRET imaging, but provide invaluable lessons on the general relationships between structure and photophysics in these fluorescent proteins. In addition to CFPs obtained from the Y66W substitution in AvGFP variants, one should not overlook the many other routes towards blue fluorescence offered by the GFP superfamily [2, 4, 57]. In particular, the Anthozoa class is rich in GFP-like proteins with blueshifted emissions [21, 58]. Of prominent interest for FRET imaging is the monomeric teal fluorescent protein mTFP1, developed from the Clavularia coral variant cFP484 [21]. Unlike CFPs, this variant carries the tyrosine-based chromophore common to all naturally occurring GFPs. Owing to a peculiar microenvironment, this chromophore exhibits a blue– green fluorescence that is more than twice as bright as mTurquoise [21]. mTFP1 is also very photostable [1, 21] and displays a near-single-exponential emission decay [24, 26], making it an attractive choice for FLIM techniques. The absorption and fluorescence spectra of


mTFP1 are slightly redshifted relative to tryptophanbased CFPs, which may require dedicated optical filters, and can increase cross-talk problems in FRET detection. Nevertheless, the use of mTFP1 may have compelling advantages if maximum fluorophore brightness is the chief concern.

4 Brighter CFPs with simplified photophysics In contrast, it is important to note that all new CFPs listed in Table 1 share very similar absorption and fluorescence spectra [51, 53, 54, 56], showing the typical double-hump shape of indole-based chromophore bands, with absorption maxima at (433 ± 2) nm and emission maxima at (473 ± 2) nm (Fig. 1). These fluorescent proteins can thus readily replace each other within the same FRET detection setup. Among these variants, gains in relative brightness (ε × φ) are mostly obtained through increases in the fluorescence quantum yield, with little change to the absorption efficiency. To the best of our knowledge, the 93% quantum yield of mTurquoise2 [55] ranks as the highest value reported today for a monomeric fluorescent protein (although fluorescence quantum yields of 0.92 and 0.95 have been reported for two naturally occurring and multimeric fluorescent proteins from copepod Pontella mimocerami [59] and cephalocordate Branchiostoma lanceolatum [29], respectively). These new CFPs also display unprecedented long fluorescence lifetimes close to 4 ns (Table 1). As expected, increases in the fluorescence quantum yield across variants are roughly paralleled by equivalent increases in their average fluorescence lifetime, although ECFP deviates somewhat from this pro-

Figure 1. Near indistinguishable absorption (thin continuous lines) and fluorescence spectra (various solid and dashed lines) of different purified CFPs, normalized at unit maximum intensity of their chromophore bands. Emission spectra were recorded at 20°C with excitation at 420 nm. Compilation of data published in [51] and [56].

183




Figure 2. (A) Marked simplification of the fluorescence emission of Aquamarine and mTurquoise, relative to ECFP: distributions of fluorescence lifetimes obtained from TCSPC fluorescence decays of the purified proteins; redrawn with permission from [56] and [45]. (B) Comparison of the fluorescence lifetime distributions of purified ECFP and ECFP-A206K [79]. Briefly, these fluorescence lifetime distributions are obtained by modeling the fluorescence decays with i 150

a generalized form of F(t)  F0

 c expt /   as the sum of a fixed set of about 150 decays with lifetimes values of τ ranging from 10 ps to 10 ns; the i

i

i

i 1

corresponding pre-exponential amplitudes, ci, are optimized parameters. In such analyses, and contrary to conventional methods of discrete exponential fitting, no a priori constraint is imposed on the number of fluorescent components, but the existence of such components is inferred from the reproducible observation of separate peaks in the final distribution, c(τ). It can be shown that the surface of each such peak equals the amplitude that would be obtained for the equivalent lifetime component in a discrete multiexponential fit of the same data [83, 84]. (C) Numerical simulations illustrating the difficulties in detecting the short lifetime components of ECFP, and their possible consequences on measurement accuracy: the successive removal of the two minor, shortest components (light grey and dark grey peaks) in the fluorescence lifetime distribution of ECFP, leads to marked increases in the apparent average lifetime  

i cii . (D) Comparison of the fluorescence decays computed before and after the removal of the two short components, showing

their very small contribution to the signal. Inset: expansion of the short time range below 1 ns.

portionality relationship. Strikingly, among a large series of CFPs, we found that higher quantum yields and longer fluorescence lifetimes were correlated to a simplified photophysics, that is, to a more homogeneous excitedstate population [51]. This is illustrated by the distributions of fluorescence lifetimes of ECFP, Aquamarine, and mTurquoise displayed in Fig. 2A, as obtained from highresolution TCSPC fluorescence decay measurements. With minor short components of 11, 13, and 17%, respec-

184

tively, Aquamarine (Fig. 2A), mTurquoise (Fig. 2A), and Cerulean-T65S, stand in sharp contrast to the multiexponential decays of ECFP and Cerulean (48 and 36%, respectively, of short lifetimes) [45, 51, 56]. Having a cyan donor with a long, near-single-exponential emission decay is a major breakthrough for quantitative FRET imaging based on fluorescence lifetime detection. First, removal of the multiple short lifetimes of ECFP will markedly improve accuracy in monitoring the





average FRET donor lifetimes. Measurement errors arising from the occurrence of short lifetimes in fluorescence decays are illustrated in Fig. 2C and D. The complete fluorescence lifetime distribution of ECFP (Fig. 2C, dashed line), or a truncated lifetime distribution missing the shortest minor component below 0.3 ns (light gray surface of Fig. 2C), or a distribution obtained after deletion of all short components below 1 ns (light and dark grey surfaces of Fig. 2C), have been used to generate different synthetic fluorescence decays that are compared after tail-normalization in Fig. 2D. These simulations show that all experimental information related to the shortest minor lifetime is restricted to the first few hundred picoseconds of the decay. This time range is very difficult to monitor reproducibly because it is strongly influenced by the instrumental response function: defective detection in this time range may increase the apparent average lifetime from 2.44 to 2.62 ns, that is, by up to 7%. Completely defective detection in the region below 1 ns, or alternatively, tail fitting or coarse fits to single-exponential models, which is a frequent practice with time-gated FLIM setups [9, 18, 26], will lead to even larger errors. Besides improving the measurement accuracy, the use of a FRET donor with a long, single-exponential emission decay also allows more detailed, in-depth analyses of complex FRET systems. Biological samples nearly always comprise different populations of donor fluorophores, arising, for example, from mixtures of interacting and noninteracting molecules or from open and closed states of single-chain biosensors. For example, in intermolecular FRET experiments, the apparent FRET efficiency, Eapp, is frequently interpreted as the product of Eapp = fDEmax, in which Emax is the FRET efficiency within the pure bimolecular complex and fD the fraction of donors engaged in this complex [24]. In most cases, estimating fD is far more crucial than determining the exact value of Emax. Providing that the respective fluorescence lifetimes of free and bound donors, τF and τB, are sufficiently different, a biexponential analysis of the donor fluorescence decay, F(t), given by Eq. (1): F (t)  F0 cB exp  t /  B   (1  cB ) exp  t /  F  

(1)

may allow the determination of this fraction. Indeed, the normalized pre-exponential amplitudes (cB and 1–cB) associated with the two lifetimes should reflect the relative molar fractions (fD and 1–fD) of the corresponding donor species, according to Eq. ((2), which gives the preexponential amplitudes, ci, of the fluorescence decay in the case of a fluorophore mixture [60]: ci 

fi  i  i k ri  fi i i k ri

i

fi

i fi

(2)

in which εi is the molar absorption coefficient, ϕi is the detected fraction of the fluorescence spectrum, and kri is


the radiative rate of each donor species i. The last three quantities actually cancel out because they should remain similar in different FRET donor populations. This approach has been applied in a few cases to estimate fD, using single-exponential FRET donors such as EGFP, mTFP1, or mTurquoise [26, 27, 61]. The use of single-exponential donors can also substantially improve the output of other fitting or non-fitting analyses of FLIM data, such as global analysis of FLIM images [10], the phasor plots proposed by Gratton and co-workers [62], or the minimal fraction of interacting donors, mfD, proposed by Tramier and Padilla-Parra [17]. All of these quantified approaches may ultimately lead, for example, to the estimate of effective binding affinities of intracellular complexes [63]. The robust, single-exponential decays of the new CFPs will extend the feasibility of such advanced analyses to a broad range of new biological questions.

5 Improved photostability Most newly engineered CFPs reportedly display improved overall photostability [51, 53, 54, 56]. However, GFP photoreactions encompass at least three different processes, namely, photobleaching, photoconversion, and photoswitching, that need to be distinguished. Photobleaching, that is, the progressive, irreversible loss of fluorescence upon prolonged illumination, is probably the most critical aspect for many imaging applications. In Table 1, we report the photobleaching times of different cytosolic CFPs determined under identical, moderate wide-field illumination. We obtained fairly consistent results with immobilized purified proteins [51]. With regards to photobleaching, Aquamarine, Cerulean-T65S, or mTurquoise all display significantly improved photostability relative to ECFP with, in the case of mTurquoise, a lifetime under irradiation extended by a factor of two. Before they bleach, irradiated GFPs may maintain some fluorescence, yet display various, irreversible phototransformations; a process called photoconversion. Whereas EGFP and EYFP undergo major spectral changes towards red- and cyan-like forms, respectively [40, 64], irradiated ECFP mostly displays a pronounced decrease in its fluorescence lifetime [20], which indicates the formation of photoproducts with similar spectral properties, but shortened fluorescence lifetimes. In this respect, mTurquoise and Aquamarine behave very similarly to ECFP: for an equivalent degree of fluorescence loss by photobleaching, the fluorescence lifetimes of all three proteins undergo the same decrease (Fig. 3A). However, the extent of decrease of the fluorescence lifetime does not equal the extent of intensity loss (slope different from one in Fig. 3A). This lack of proportionality shows that photoconversion and photobleaching of these proteins involve distinct fluorescent and non-fluorescent photoproducts,

185


Biotechnol. J. 2014, 9, 180–191



Figure 3. Irreversible and reversible photoreactions of CFPs. (A) Irreversible light-induced conversion of CFPs to photoproducts with shorter fluorescence lifetimes: relationship between the observed intensity loss through photobleaching and the simultaneous decrease in the measured average lifetime. Data was obtained by FLIM analyses under continuous wide-field illumination at 0.2 W/cm2 of the cytosolic proteins expressed in MDCK cells; redrawn with permission from [56]. (B) Schematic view of the photoisomerization properties of the isolated CFP chromophore for an example of a simple torsion along the τ dihedral angle (isomer nomenclature according to Voliani et al. [68]): light absorption by the planar cis or trans ground-state isomers leads to a similar excited-state S1, in which the two aromatic moieties adopt a preferred perpendicular configuration. kf and k’f represent the de-excitation paths back to the cis and trans groundstate isomers, respectively, whereas k– and k+ represent their slow thermal equilibration. (C) Reversible photoswitching responses of purified CFPs immobilized on agarose beads. After prior equilibration in the dark, sudden wide-field lamp illumination through a CFP filter set at 0.2 W/cm2 was applied for 10 s. The return of fluorescence after 3 min in the dark was checked by taking a few snapshots. Continuous lines are best fits to a reversible two-state model; redrawn with permission from [51].

respectively. CFP photoconversion can take place within a few seconds of high-power illumination and should always be very carefully controlled because it may dangerously corrupt FLIM–FRET measurements [20, 65]. Finally, electronically excited GFPs also undergo transient conversions to new dark or emissive species that are fully and slowly reversed in the dark or under illumination at specific wavelengths; a phenomenon now commonly referred to as reversible photoswitching [57, 66, 67]. Model isolated GFP chromophores display efficient reversible photoisomerization properties between different cis and trans diastereoisomers [68] (Fig. 3B). Because these chromophore isomers have strongly overlapping absorption spectra, the photoreaction takes place in both directions. The system equilibrates only slowly in the dark due to the large isomerization barrier in the ground state. Despite steric hindrance, chromophore photoisomerization seems to occur easily inside well-folded fluorescent proteins, as demonstrated by the X-ray crystallographic structures of many, so-called reversibly switchable fluorescent proteins

186

(RSFPs) [67]. Photoisomerization quantum efficiencies are typically 10–20% for the isolated chromophores [68], whereas photoswitching efficiencies range from 10–4 to 15% for different RSFPs [67]. Inside fluorescent proteins, chromophore isomerization may be accompanied by other specific perturbations, such as changes in the protonation state of tyrosine-based chromophores, leading to large spectral shifts in their absorption bands [66]. Other photoreactions, such as excited-state proton transfer (ESPT), are frequently coupled to chromophore photoisomerization [57, 67]. ESPT is sometimes proposed to be the driving force in GFP photoswitching [69, 70]. Finally, a peculiar type of reversible photochemical reaction was recently reported in Dreiklang, which is a photoswitchable Citrine variant. In this case, photoswitching involves the reversible hydration of the chromophore imidazolinone moiety; hydration is reversed by illumination of the photoproduct absorption band at 340 nm [71]. It was only recognized recently that CFP proteins were very efficient RSFPs [39, 51, 72]. The photoswitching of



Biotechnol. J. 2014, 9, 180–191


ECFP is characterized by a rapid partial loss of fluorescence observed at very low illumination power, whereas Cerulean displays an even more pronounced response (Fig. 3C). Fluorescence then fully recovers in the dark on the timescale of minutes, and the repeatability of the experiment is only limited by the slow concomitant photobleaching process [51]. At moderate irradiance, we showed that the transients could be described by a simple reversible model, involving two photoactivated “on” and “off” reactions taking place simultaneously. The high quantum efficiency of both photoreactions (Φon–off =1% and Φoff–on=6% for ECFP) [51], and the fact that they are activated at the same excitation wavelength, strongly suggest a photoisomerization reaction, as depicted in Fig. 3B, although this idea still requires further experimental support. Due to the slow timescale of recovery, the photoswitching properties of ECFP and Cerulean are likely to impede the stable monitoring of ratiometric FRET traces and more generally fluorescence quantifications. We found that the T65S mutation alone considerably dampened the photoswitching amplitude (Fig. 3C), due to a 10-fold decrease if the on–off reaction rate [51]. Aquamarine and mTurquoise display barely detectable transients (Fig. 3C) that result from 40- and 100-fold, respectively, decreases in the same reaction rate, as compared with ECFP [51]. Using Aquamarine as the FRET donor in the AKAR2.2 biosensor contributed to a marked stabilization of its ratiometric traces as a function of the imaging frequency [56].

6 Reduced environmental sensitivity Sensitivity to pH is a major issue in the use of GFPs and YFPs, which complicates any reliable bioanalytical sensing in the many compartments involved in cell bioenergetics, secretion, and transport [1, 2]. Notably, in the acid pH range, most popular GFP variants display a total loss in fluorescence emission with apparent pK values between 5 and 7, that is, very close to physiological pH. In GFPs with tyrosine-based chromophores, this drop in fluorescence is paralleled by an equivalent loss in absorbance of the anionic chromophore, and is thus ascribed to the pH-dependent protonation of the chromophore phenol. However, a similar loss of fluorescence, with only small changes in absorbance, is also observed at acid pH values in ECFP, the chromophore of which does not bear a protonatable group [1, 21]. This fluorescence drop is accompanied by a concomitant decrease in fluorescent lifetime [45]. We estimated a half-transition point at pH 5.6 for the fluorescence intensity of purified ECFP in solution [51], and at pH 6 for its fluorescence lifetime inside secretory granules of PC12 cells [73]. The ECFP lifetime is thus well suited for quantitative FLIM-based pH determinations of acidic organelles [73]. However, when ECFP is used as a FRET donor, this pH sensitivity is



potentially a major flaw. Strikingly, all new CFPs display an outstanding stability towards acid pH, with pK1/2 around or below 3.5 (Table 1); a performance challenged today by very few other GFP variants [1, 2, 29]. A strong contribution to signal variability is also likely to be associated with the pronounced temperature sensitivity of ECFP. Indeed, its fluorescence intensity and lifetime both decrease by about 2% per 1°C when the temperature increases in the 20 to 40°C range [45]. By contrast, new generations of CFPs, such as Aquamarine and mTurquoise, display a markedly reduced sensitivity to temperature, with decreases of only 0.5% per 1°C. We also found that the intracellular fluorescent lifetime of Aquamarine was remarkably insensitive to genetic fusion or to the subcellular location [56]: for membrane-targeted MyrPalm-Aquamarine, we observed a fluorescence lifetime of 3.73 ns, that is, decreased by only 6% relative to its cytosolic value, whereas the lifetime of Myr-Palm-ECFP decreased by 36%. This environmental insensitivity of Aquamarine is particularly advantageous for the design of reliable FRET “donor-only” samples and for improving the general accuracy of FRET measurements. We have hypothesized that a reduced environmental sensitivity, together with major gains in fluorescence quantum yield, simplified photophysics, and apparent blockage of photoswitching, are all different consequences of a significant rigidification of the chromophore in Aquamarine; an idea that probably also holds for the other new CFPs [51, 56]. Compared with less-optimized CFPs, mTurquoise and Aquamarine both suffer from a delayed fluorescence growth when expressed in bacterial cultures, as well as from a delayed recovery of fluorescence along refolding experiments in vitro [55, 56]. These slow kinetics are probably the price to pay for having more compact, rigid proteins. Nevertheless, we met no difficulty in obtaining high fluorescence levels in any type of mammal cell line expressing Aquamarine or mTurquoise, as early as one day after transfection. Also, the average brightness of these cytosolic proteins is increased roughly by a factor of two relative to ECFP [53, 56]. This value, in agreement with their relative quantum yields measured in vitro, shows that these fluorescent proteins fold and mature efficiently in eukaryotic cells.

7 Structural bases of CFP ameliorations Aquamarine is obtained by only two mutations of ECFP (T65S, H148G), yet ranks among the best CFP variants. This brings worthwhile information on the structural basis of their improved properties. First, because all other highly optimized CFPs carry mutations at the same two positions, 148 and 65 (Table 1), it is likely that their improved photophysics arises chiefly from these two mutations. Besides these mutations, only two other muta-

187


Biotechnol. J. 2014, 9, 180–191


tions, S72A and A206K, are common to mTurquoise, mTurquoise2, and mCerulean3. The monomerizing A206K mutation is expected to forbid weak dimer formation in the AvGFP variants [74]. In agreement with previous reports, we observed no significant impact of this mutation on the ECFP photophysics, as demonstrated, for example, by the undistinguishable fluorescence lifetime distributions of ECFP and ECFPA206K (Fig. 2B). Dimerization between CFPs and YFPs increases the FRET levels and is sometimes favored to improve the dynamics of biosensor responses [75–77]. Nevertheless, introducing the A206K mutation remains a safe measure if fluorescent protein dimerization must be strictly avoided. Notably, the A206K mutation has no detectable incidence on the intracellular interactions of free cytosolic ECFP and EYFP [18], and does not change the apparent dissociation constant of the fluorescenttagged ERα receptor [78]. Contrary to EYFP, the purified ECFP protein does not form homodimers due to the I146N mutation, which modifies its dimerization interface [79]. With this view, it might be advisable to evaluate the impact of the N146F mutation of mTurquoise2 on its dimerization properties. The T65S mutation corresponds to the simple removal of the extra methyl group of a threonine, which restores the wild-type serine 65 residue of AvGFP. This conservative mutation seems to have beneficial effects in many different contexts, including not only CFPs, but also blue fluorescent proteins and GFPs (reviewed in [51]). The single-point mutation T65S results in a 50% increase in the fluorescence quantum yield of SCFP3A [53], a 45% increase in that of mCerulean2 [54], and 48 and 25% increases in those of ECFP and Cerulean, respectively. Its strong effect on the CFP photoswitching properties suggests that residue 65 has a major role in controlling the excited-state chromophore torsions. However, X-ray crystallographic studies of SCFP3A and mTurquoise [55] have revealed extremely similar, if not identical, local 3D structures, with a root-mean-square (RMS) distance of 0.08 Å between all heavy atoms of the chromophore cavity. The only notable difference is a small displacement of the Leu 220 aliphatic side chain, due to steric repulsion by the extra methyl group of Thr 65. However, because this side chain is relatively flexible and not in direct contact with the chromophore system, it is hard to understand how it might control the excited-state chromophore torsions so drastically. Mutations at position 220 actually resulted in only mild perturbations of the chromophore photophysics [55]. In other words, there seems to be no clear structural correlation to the extensive photophysical consequences of the T65S mutation in mTurquoise. Based on the comparative analyses of chromophore cavities of X-ray crystallographic structures of different AvGFP variants, we identified a conformational mechanism that might account for this puzzling situation [51]. We found that, depending on the protein and crystallo-

188


Figure 4. Changes in hydrogen-bonding connectivity and structural water organization associated with the Up and Down conformations of the residue 65 hydroxyl in CFPs: example of the ECFP structures 2WSN (solid) [81] , and 1OXD (transparent) [48]; redrawn with permission from [56] .

graphic structure, residue 65 adopted two different “Up” and “Down” orientations of its hydroxyl group, as depicted in Fig. 4 for ECFP. First, all green GFPs carrying a wildtype serine in position 65 adopt the Down configuration, while all those carrying a threonine display instead the Up configuration. Second, in CFPs carrying a threonine, such as ECFP and Cerulean, both configurations can be observed, depending on the crystallographic structure, which suggests a higher degree of conformational freedom in these proteins [48, 80, 81]. In GFPs as well as CFPs, the residue 65 hydroxyl in the Down configuration connects to a well-ordered, conserved water layer in direct contact with the chromophore (Fig. 4). When the residue 65 hydroxyl adopts the Up configuration, it disconnects from these water molecules and makes two new hydrogen bonds: one with the imidazolinone nitrogen of the chromophore and the other with the valine 61 mainchain carbonyl. In ECFP and Cerulean, the water layer then becomes clearly disorganized, resulting in looser packing of the chromophore. If a conformational equilibrium takes place between significant populations of the Up and Down configurations in CFPs, any displacement of this equilibrium (e.g. as possibly favored by the T65S mutation) may result in marked changes in the average chromophore photophysics in solution. The crystallogenesis process, by selecting a single, most stable conformation, would lose track of this dynamic equilibrium. However, specific crystal growth conditions, such as acid pH in the case of Cerulean [80], may in some cases favor the alternate conformation.





Two very different substitutions, D and G, are observed at position 148 in the best CFPs, making a structural interpretation less clear at the moment. We found that, in general, amino acid residues with small side chains performed best at position 148 in ECFP [56], which suggests that they simply provide sufficient room to accommodate the bulky tryptophan-based chromophore without further energetic strain on the rest of the protein. It is also worth noting that exchanging the mutation H148G to H148D in Aquamarine substantially decreases its pH stability [56]. Therefore, the high pH stability of mTurquoise or Cerulean-T65S, both carrying an aspartate at position 148, requires the contribution of other mutations specific to these proteins. Indeed, we found that the S72A mutation, common to the two proteins, decreases the pH1/2 of ECFP by 0.4 units [82].

Fabienne Mérola has academic training in chemical physics and biology, and is an expert in molecular fluorescence spectroscopy. After post-doctoral studies at the Medical Biophysics Department of the Karolinska Institute (Stockholm), she was appointed as a synchrotron beam-line scientist at LURE, Orsay, where she studied the biophysical chemistry of various enzymes and receptors. She now works at the LCP on the development of new quantitative probes and methods for in situ biochemical imaging of living systems. Her team carries out advanced FRET and FLIM microscopy studies, and combines photophysical and molecular modeling approaches for the design of new fluorescent protein biosensors.

8 Conclusions and perspectives

9 References

We now have at hand several bright CFPs with robust, environmentally insensitive, single-exponential emission decays, which opens up new avenues for many imaging applications. Owing to the stepwise development of Aquamarine, a simple method, based on widely accessible site-directed mutagenesis kits, can be proposed to upgrade the many current gene constructs carrying ECFP or Cerulean to access similarly improved performances. Fluorescence quantum yields approaching unity also indicate that the level of non-radiative processes competing with fluorescence emission has become negligible. Further major gains in brightness can now only be obtained by increasing the absorption efficiency, which will probably require changing the electronic conjugated system. These photophysical performances are thus presumably close to being ultimate, for fluorescent proteins carrying a tryptophan-based chromophore. Nevertheless, the detailed comparison of Aquamarine and other CFP variants brings many valuable insights into the structural determinants of GFP fluorescence. These clues will be very useful, not only to inspire the design of new, improved fluorescent proteins, but also to guide the rational engineering of new functions, such as direct specific sensing or optogenetic properties, into these proteins. Other exciting challenges remain ahead, for example, finding routes to improve further their photostability and to progress towards accurate, routine FRET quantifications within living cells.

[1] Shaner, N. C., Patterson, G. H., Davidson, M. W., Advances in fluorescent protein technology. J. Cell Sci. 2007, 120, 4247–4260. [2] Chudakov, D. M., Matz, M. V., Lukyanov, S., Lukyanov, K. A., Fluorescent proteins and their applications in imaging living cells and tissues. Physiol. Rev. 2010, 90, 1103–1163. [3] Balla, T., Green light to illuminate signal transduction events. Trends Cell Biol. 2009, 19, 575–586. [4] Newman, R. H., Fosbrink, M. D., Zhang, J., Genetically encodable fluorescent biosensors for tracking signaling dynamics in living cells. Chem Rev. 2011, 111, 3614–3666. [5] Aoki, K., Komatsu, N., Hirata, E., Kamioka, Y. et al., Stable expression of FRET biosensors: A new light in cancer research. Cancer Sci. 2012, 2011, 1349–7006. [6] Perron, A., Akemann, W., Mutoh, H., Knopfel, T., Genetically encoded probes for optical imaging of brain electrical activity. Prog. Brain Res. 2012, 196, 63–77. [7] Morris, M. C., Fluorescent biosensors-probing protein kinase function in cancer and drug discovery. Biochim. Biophys. Acta 2013, 1834, 1387–1395. [8] Valeur, B., Molecular Fluorescence. Principles and Applications, Wiley-VCH, Weinheim, 2001. [9] Sipieter, F., Vandame, P., Spriet, C., Leray, A. et al., From FRET imaging to practical methodology for kinase activity sensing in living cells. Prog. Mol. Biol. Transl. Sci. 2013, 113, 145–216. [10] Grecco, H. E., Roda-Navarro, P., Girod, A., Hou, J. et al., In situ analysis of tyrosine phosphorylation networks by FLIM on cell arrays. Nat. Methods 2010, 7, 467–472. [11] Hoppe, A. D., Swanson, J. A., Cdc42, Rac1, and Rac2 display distinct patterns of activation during phagocytosis. Mol. Biol. Cell 2004, 15, 3509–3519. [12] Adjobo-Hermans, M. J., Goedhart, J., van Weeren, L., Nijmeijer, S. et al., Real-time visualization of heterotrimeric G protein Gq activation in living cells. BMC Biol. 2011, 9, 1741–7007. [13] Gervasi, N., Hepp, R., Tricoire, L., Zhang, J. et al., Dynamics of protein kinase A signaling at the membrane, in the cytosol, and in the nucleus of neurons in mouse brain slices. J. Neurosci. 2007, 27, 2744–2750. [14] Miyawaki, A., Development of probes for cellular functions using fluorescent proteins and fluorescence resonance energy transfer. Annu. Rev. Biochem. 2011, 80, 357–373.

A.F., D.B.B., and C.Z. acknowledge doctoral grants from the French Ministry of Superior Education and Research, the Région Ile-de-France (C’Nano IdF), and the Interdisplinary Initiative of IDEX campus Paris-Saclay, respectively. We also acknowledge support from Nikon France, CNRS, Université Paris Sud, and ANR. The authors declare no conflict of interest.


189


Biotechnol. J. 2014, 9, 180–191


[15] Subach, F. V., Zhang, L., Gadella, T. W., Gurskaya, N. G. et al., Red fluorescent protein with reversibly photoswitchable absorbance for photochromic FRET. Chem. Biol. 2010, 17, 745–755. [16] Bizzarri, R., Serresi, M., Cardarelli, F., Abbruzzetti, S. et al., Single amino acid replacement makes Aequorea victoria fluorescent proteins reversibly photoswitchable. J. Am. Chem. Soc. 2010, 132, 85–95. [17] Padilla-Parra, S., Tramier, M., FRET microscopy in the living cell: Different approaches, strengths and weaknesses. Bioessays 2012, 34, 369–376. [18] Grailhe, R., Merola, F., Ridard, J., Couvignou, S. et al., Monitoring protein interactions in the living cell through the fluorescence decays of the cyan fluorescent protein. ChemPhysChem 2006, 7, 1442–1454. [19] Wolber, P. K., Hudson, B. S., An analytic solution to the Förster energy transfer problem in two dimensions. Biophys. J. 1979, 28, 197– 210. [20] Tramier, M., Zahid, M., Mevel, J. C., Masse, M. J. et al., Sensitivity of CFP/YFP and GFP/mCherry pairs to donor photobleaching on FRET determination by fluorescence lifetime imaging microscopy in living cells. Microsc. Res. Tech. 2006, 69, 933–939. [21] Ai, H. W., Henderson, J. N., Remington, S. J., Campbell, R. E., Directed evolution of a monomeric, bright and photostable version of Clavularia cyan fluorescent protein: Structural characterization and applications in fluorescence imaging. Biochem. J. 2006, 400, 531– 540. [22] Goedhart, J., Vermeer, J. E., Adjobo-Hermans, M. J., van Weeren, L. et al., Sensitive detection of p65 homodimers using red-shifted and fluorescent protein-based FRET couples. PLoS One 2007, 2, e1011. [23] Ai, H. W., Hazelwood, K. L., Davidson, M. W., Campbell, R. E., Fluorescent protein FRET pairs for ratiometric imaging of dual biosensors. Nat. Methods 2008, 5, 401–403. [24] Padilla-Parra, S., Auduge, N., Lalucque, H., Mevel, J. C. et al., Quantitative comparison of different fluorescent protein couples for fast FRET-FLIM acquisition. Biophys. J. 2009, 97, 2368–2376. [25] Subach, O. M., Cranfill, P. J., Davidson, M. W., Verkhusha, V. V., An enhanced monomeric blue fluorescent protein with the high chemical stability of the chromophore. PLoS One 2011, 6, e28674. [26] Walther, K. A., Papke, B., Sinn, M. B., Michel, K. et al., Precise measurement of protein interacting fractions with fluorescence lifetime imaging microscopy. Mol. Biosyst. 2011, 7, 322–336. [27] Laine, R., Stuckey, D. W., Manning, H., Warren, S. C. et al., Fluorescence lifetime readouts of Troponin-C-based calcium FRET sensors: a quantitative comparison of CFP and mTFP1 as donor fluorophores. PLoS One 2012, 7, e49200. [28] Lam, A. J., St-Pierre, F., Gong, Y., Marshall, J. D. et al., Improving FRET dynamic range with bright green and red fluorescent proteins. Nat. Methods 2012, 9, 1005–1012. [29] Shaner, N. C., Lambert, G. G., Chammas, A., Ni, Y. et al., A bright monomeric green fluorescent protein derived from Branchiostoma lanceolatum. Nat. Methods 2013, 10, 407–409. [30] Tsien, R. Y., The green fluorescent protein. Annu. Rev. Biochem. 1998, 67, 509–544. [31] Kardash, E., Bandemer, J., Raz, E., Imaging protein activity in live embryos using fluorescence resonance energy transfer biosensors. Nat. Protoc. 2011, 6, 1835–1846. [32] Kamioka, Y., Sumiyama, K., Mizuno, R., Sakai, Y. et al., Live imaging of protein kinase activities in transgenic mice expressing FRET biosensors. Cell 2012, 37, 65–73. [33] Rinnenthal, J. L., Bornchen, C., Radbruch, H., Andresen, V. et al., Parallelized TCSPC for dynamic intravital fluorescence lifetime imaging: Quantifying neuronal dysfunction in neuroinflammation. PLoS One 2013, 8, e60100.

190


[34] Nobis, M., McGhee, E. J., Morton, J. P., Schwarz, J. P. et al., Intravital FLIM-FRET imaging reveals dasatinibinduced spatial control of Src in pancreatic cancer. Cancer Res. 2013, 73, 4674–4686. [35] Aye-Han, N. N., Allen, M. D., Ni, Q., Zhang, J., Parallel tracking of cAMP and PKA signaling dynamics in living cells with FRET-based fluorescent biosensors. Mol. Biosyst. 2012, 8, 1435–1440. [36] Kominami, K., Nagai, T., Sawasaki, T., Tsujimura, Y. et al., In vivo imaging of hierarchical spatiotemporal activation of caspase-8 during apoptosis. PLoS One 2012, 7, e50218. [37] Griesbeck, O., Baird, G. S., Campbell, R. E., Zacharias, D. A. et al., Reducing the environmental sensitivity of yellow fluorescent protein. Mechanism and applications. J Biol. Chem. 2001, 276, 29188– 29194. [38] Tramier, M., Gautier, I., Piolot, T., Ravalet, S. et al., Picosecond-hetero-FRET microscopy to probe protein–protein interactions in live cells. Biophys. J. 2002, 83, 3570–3577. [39] Sinnecker, D., Voigt, P., Hellwig, N., Schaefer, M., Reversible photobleaching of enhanced green fluorescent proteins. Biochemistry 2005, 44, 7085–7094. [40] Valentin, G., Verheggen, C., Piolot, T., Neel, H. et al., Photoconversion of YFP into a CFP-like species during acceptor photobleaching FRET experiments. Nat. Methods 2005, 2, 801. [41] Willemse, M., Janssen, E., de Lange, F., Wieringa, B. et al., ATP and FRETÑa cautionary note. Nat. Biotechnol. 2007, 25, 170–172. [42] Koushik, S. V., Vogel, S. S., Energy migration alters the fluorescence lifetime of Cerulean: Implications for fluorescence lifetime imaging Förster resonance energy transfer measurements. J. Biomed. Opt. 2008, 13, 031204. [43] Heim, R., Prasher, D. C., Tsien, R. Y., Wavelength mutations and posttranslational autoxidation of green fluorescent protein. Proc. Natl. Acad. Sci. USA 1994, 91, 12501–12504. [44] Heim, R., Tsien, R. Y., Engineering green fluorescent protein for improved brightness, longer wavelengths and fluorescence resonance energy transfer. Curr. Biol. 1996, 6, 178–182. [45] Villoing, A., Ridhoir, M., Cinquin, B., Erard, M. et al., Complex fluorescence of the cyan fluorescent protein: Comparisons with the H148D variant and consequences for quantitative cell imaging. Biochemistry 2008, 47, 12483–12492. [46] Borst, J. W., Willemse, M., Slijkhuis, R., van der Krogt, G. et al., ATP changes the fluorescence lifetime of cyan fluorescent protein via an interaction with His148. PLoS One 2010, 5, e13862. [47] Seifert, M. H., Ksiazek, D., Azim, M. K., Smialowski, P. et al., Slow exchange in the chromophore of a green fluorescent protein variant. J. Am. Chem. Soc. 2002, 124, 7932–7942. [48] Bae, J. H., Rubini, M., Jung, G., Wiegand, G. et al., Expansion of the genetic code enables design of a novel “gold” class of green fluorescent proteins. J. Mol. Biol. 2003, 328, 1071–1081. [49] Rizzo, M. A., Springer, G. H., Granada, B., Piston, D. W., An improved cyan fluorescent protein variant useful for FRET. Nat. Biotechnol. 2004, 22, 445–449. [50] Kremers, G. J., Goedhart, J., van Munster, E. B., Gadella, T. W. J., Cyan and yellow super fluorescent proteins with improved brightness, protein folding, and FRET Förster radius. Biochemistry 2006, 45, 6570–6580. [51] Fredj, A., Pasquier, H., Demachy, I., Jonasson, G. et al., The single T65S mutation generates brighter cyan fluorescent proteins with increased photostability and pH insensitivity. PLOS One 2012, 7, e49149. [52] Nagai, T., Ibata, K., Park, E. S., Kubota, M. et al., A variant of yellow fluorescent protein with fast and efficient maturation for cell-biological applications. Nat. Biotechnol. 2002, 20, 87–90. [53] Goedhart, J., van Weeren, L., Hink, M. A., Vischer, N. O. et al., Bright cyan fluorescent protein variants identified by fluorescence lifetime screening. Nat. Methods 2010, 7, 137–139.



Biotechnol. J. 2014, 9, 180–191


[54] Markwardt, M. L., Kremers, G. J., Kraft, C. A., Ray, K. et al., An improved cerulean fluorescent protein with enhanced brightness and reduced reversible photoswitching. PLoS One 2011, 6, e17896. [55] Goedhart, J., von Stetten, D., Noirclerc-Savoye, M., Lelimousin, M. et al., Structure-guided evolution of cyan fluorescent proteins towards a quantum yield of 93%. Nat. Commun. 2012, 3, doi: 10.1038. [56] Erard, M., Fredj, A., Pasquier, H., Betolngar, B. D. et al., Minimum set of mutations needed to optimize cyan fluorescent proteins for live cell imaging. Mol. Biosyst. 2013, 8, 258–267. [57] Merola, F., Levy, B., Demachy, I., Pasquier, H., Photophysics and Spectroscopy of Fluorophores in the Green Fluorescent Protein Family. Springer Series on Fluorescence. 2010, 8, 347–383. [58] Matz, M. V., Marshall, N. J., Vorobyev, M., Are corals colorful? Photochem. Photobiol. 2006, 82, 345–350. [59] Hunt, M. E., Scherrer, M. P., Ferrari, F. D., Matz, M. V., Very bright green fluorescent proteins from the Pontellid copepod Pontella mimocerami. PLoS One 2010, 5, 0011517. [60] Mérola, F., State Thesis in Life Sciences, Université Paris XI, 1995. [61] Verveer, P. J., Wouters, F. S., Reynolds, A. R., Bastiaens, P. I., Quantitative imaging of lateral ErbB1 receptor signal propagation in the plasma membrane. Science 2000, 290, 1567–1570. [62] Digman, M. A., Caiolfa, V. R., Zamai, M., Gratton, E., The phasor approach to fluorescence lifetime imaging analysis. Biophys. J. 2008, 94, L14–L16. [63] Cardarelli, F., Bizzarri, R., Serresi, M., Albertazzi, L. et al., Probing nuclear localization signal-importin alpha binding equilibria in living cells. J. Biol. Chem. 2009, 284, 36638–36646. [64] Bogdanov, A. M., Mishin, A. S., Yampolsky, I. V., Belousov, V. V. et al., Green fluorescent proteins are light-induced electron donors. Nat. Chem. Biol. 2009, 5, 459–461. [65] Hoffmann, B., Zimmer, T., Klocker, N., Kelbauskas, L. et al., Prolonged irradiation of enhanced cyan fluorescent protein or Cerulean can invalidate Förster resonance energy transfer measurements. J. Biomed. Opt. 2008, 13, 031205. [66] Abbruzzetti, S., Bizzarri, R., Luin, S., Nifosi, R. et al., Photoswitching of E222Q GFP mutants: “Concerted” mechanism of chromophore isomerization and protonation. Photochem. Photobiol. Sci. 2010, 9, 1307–1319. [67] Bourgeois, D., Adam, V., Reversible photoswitching in fluorescent proteins: A mechanistic view. IUBMB Life 2012, 64, 482–491. [68] Voliani, V., Bizzarri, R., Nifosi, R., Abbruzzetti, S. et al., Cis–trans photoisomerization of fluorescent-protein chromophores. J. Phys. Chem. B 2008, 112, 10714–10722. [69] Mizuno, H., Mal, T. K., Walchli, M., Kikuchi, A. et al., Light-dependent regulation of structural flexibility in a photochromic fluorescent protein. Proc. Natl. Acad. Sci. USA 2008, 105, 9227–9232. [70] Faro, A. R., Adam, V., Carpentier, P., Darnault, C. et al., Low-temperature switching by photoinduced protonation in photochromic fluorescent proteins. Photochem. Photobiol. Sci. 2010, 9, 254–262.



[71] Brakemann, T., Stiel, A. C., Weber, G., Andresen, M. et al., A reversibly photoswitchable GFP-like protein with fluorescence excitation decoupled from switching. Nat. Biotechnol. 2011, 29, 942– 947. [72] Shaner, N. C., Lin, M. Z., McKeown, M. R., Steinbach, P. A. et al., Improving the photostability of bright monomeric orange and red fluorescent proteins. Nat. Methods 2008, 5, 545–551. [73] Poea-Guyon, S., Pasquier, H., Merola, F., Morel, N. et al., The enhanced cyan fluorescent protein: A sensitive pH sensor for fluorescence lifetime imaging. Anal. Bioanal. Chem. 2013, 405, 3983–3987. [74] Zacharias, D. A., Violin, J. D., Newton, A. C., Tsien, R. Y., Partitioning of lipid-modified monomeric GFPs into membrane microdomains of live cells. Science 2002, 296, 913–916. [75] Nguyen, A. W., Daugherty, P. S., Evolutionary optimization of fluorescent proteins for intracellular FRET. Nat. Biotechnol. 2005, 23, 355–360. [76] Vinkenborg, J. L., Evers, T. H., Reulen, S. W., Meijer, E. W. et al., Enhanced sensitivity of FRET-based protease sensors by redesign of the GFP dimerization interface. ChemBioChem 2007, 8, 1119–1121. [77] Kotera, I., Iwasaki, T., Imamura, H., Noji, H. et al., Reversible dimerization of Aequorea victoria fluorescent proteins increases the dynamic range of FRET-based indicators. ACS Chem. Biol. 2010, 5, 215–222. [78] Kofoed, E. M., Guerbadot, M., Schaufele, F., Dimerization between aequorea fluorescent proteins does not affect interaction between tagged estrogen receptors in living cells. J. Biomed. Opt. 2008, 13, 031207. [79] Espagne, A., Erard, M., Madiona, K., Derrien, V. et al., Cyan fluorescent protein carries a constitutive mutation that prevents its dimerization. Biochemistry 2011, 50, 437–439. [80] Malo, G. D., Pouwels, L. J., Wang, M., Weichsel, A. et al., X-ray structure of Cerulean GFP: A tryptophan-based chromophore useful for fluorescence lifetime imaging. Biochemistry 2007, 46, 9865–9873. [81] Lelimousin, M., Noirclerc-Savoye, M., Lazareno-Saez, C., Paetzold, B. et al., Intrinsic dynamics in ECFP and Cerulean control fluorescence quantum yield. Biochemistry 2009, 48, 10038–10046. [82] Fredj, A., Doctorate Thesis, Chemistry Doctoral School, Université Paris Sud, 2012. [83] Merola, F., Rigler, R., Holmgren, A., Brochon, J. C., Picosecond tryptophan fluorescence of thioredoxin: Evidence for discrete species in slow exchange. Biochemistry 1989, 28, 3383–3398. [84] Esposito, R., Altucci, C., Velotta, R., Analysis of simulated fluorescence intensities decays by a new maximum entropy method algorithm. J. Fluoresc. 2013, 23, 203–211.

191

ISSN 1860-6768 · BJIOAM 9 (2) 171–310 (2014) · Vol. 9 · February 2014

Systems & Synthetic Biology · Nanobiotech · Medicine

2/2014 FRET imaging Synthetic probes Live-cell imaging

Fluorescent Biosensors www.biotechnology-journal.com

The Fluorescent Biosensor special issue of Biotechnology Journal is edited by Dr. May Morris and Prof. Marc Blondel. The cover image is an artistic interpretation of how fluorescent biosensors function as molecular beacons for scientists navigating a sea of molecules. Image courtesy of L. Divita and R. Wintergerst.

Biotechnology Journal – list of articles published in the February 2014 issue. Editorial: Fluorescent biosensors May C. Morris and Marc Blondel http://dx.doi.org/10.1002/biot.201400008 Review Newly engineered cyan fluorescent proteins with enhanced performances for live cell FRET imaging

Review Fluorescent biosensors for high throughput screening of protein kinase inhibitors Camille Prével, Morgan Pellerano, Thi Nhu Ngoc Van and May C. Morris

http://dx.doi.org/10.1002/biot.201300196

Fabienne Mérola, Asma Fredj, Dahdjim-Benoît Betolngar, Cornelia Ziegler, Marie Erard and Hélène Pasquier

Review FRET-based and other fluorescent proteinase probes


Hai-Yu Hu, Stefanie Gehrig, Gregor Reither, Devaraj Subramanian, Marcus A. Mall, Oliver Plettenburg and Carsten Schultz

Review Decoding spatial and temporal features of neuronal cAMP/PKA signaling with FRET biosensors Liliana R. V. Castro, Elvire Guiot, Marina Polito, Danièle Paupardin-Tritsch and Pierre Vincent

http://dx.doi.org/10.1002/biot.201300202 Review Imaging early signaling events in T lymphocytes with fluorescent biosensors Clotilde Randriamampita and Annemarie C. Lellouch

http://dx.doi.org/10.1002/biot.201300195 Review Deciphering the spatio-temporal regulation of entry and progression through mitosis Lilia Gheghiani and Olivier Gavet

http://dx.doi.org/10.1002/biot.201300194 Review Shining light on cell death processes – a novel biosensor for necroptosis, a newly described cell death program François Sipieter, Maria Ladik, Peter Vandenabeele and Franck Riquet

http://dx.doi.org/10.1002/biot.201300201 Review Genetically encoded reactive oxygen species (ROS) and redox indicators Sandrine Pouvreau

http://dx.doi.org/10.1002/biot.201300199 Technical Report Time-resolved microfluorimetry: An alternative method for free radical and metabolic rate detection in microalgae Amadine Bijoux and Anne-Cécile Ribou

http://dx.doi.org/10.1002/biot.201300217 Research Article Acri-2,7-Py, a bright red-emitting DNA probe identified through screening of a distyryl dye library Delphine Naud-Martin, Xavier Martin-Benlloch, Florent Poyer, Florence Mahuteau-Betzer and Marie-Paule Teulade-Fichou


http://dx.doi.org/10.1002/biot.201300200 Review Advances in lanthanide-based luminescent peptide probes for monitoring the activity of kinase and phosphatase Elena Pazos and M. Eugenio Vázquez




pH sensitivity of FRET reporters based on cyan and yellow fluorescent proteins.

Fluorescent vinblastine probes for live cell imaging.

Characterization of Fluorescent Proteins for Three- and Four-Color Live-Cell Imaging in S. cerevisiae.

Engineering and characterizing monomeric fluorescent proteins for live-cell imaging applications.

Self-assembled fluorescent organic nanoparticles for live-cell imaging.

A high speed multifocal multiphoton fluorescence lifetime imaging microscope for live-cell FRET imaging.

Enhanced live cell imaging via photonic crystal enhanced fluorescence microscopy.

PII Protein-Derived FRET Sensors for Quantification and Live-Cell Imaging of 2-Oxoglutarate.

Engineered fluorescent proteins illuminate the bacterial periplasm.

Fluorescent protein vectors for pancreatic islet cell identification in live-cell imaging.

Multiplexed 3D FRET imaging in deep tissue of live embryos.

Fluorescent proteins as genetically encoded FRET biosensors in life sciences.

A novel FRET 'off-on' fluorescent probe for the selective detection of Fe³⁺, Al³⁺ and Cr³⁺ ions: its ultrafast energy transfer kinetics and application in live cell imaging.

Highly fluorescent and bioresorbable polymeric nanoparticles with enhanced photostability for cell imaging.

Red fluorescent luminogen from pyrrole derivatives with aggregation-enhanced emission for cell membrane imaging.

Toward live-cell imaging of dopamine neurotransmission with fluorescent neurotransmitter analogues.

Indole-based cyanine as a nuclear RNA-selective two-photon fluorescent probe for live cell imaging.

Development of background-free tame fluorescent probes for intracellular live cell imaging.

Hoechst tagging: a modular strategy to design synthetic fluorescent probes for live-cell nucleus imaging.

Fluorescent kapakahines serve as non-toxic probes for live cell Golgi imaging.

RNA-Based Fluorescent Biosensors for Live Cell Imaging of Second Messenger Cyclic di-AMP.

A codon-optimized green fluorescent protein for live cell imaging in Zymoseptoria tritici.

Evaluation of chemical fluorescent dyes as a protein conjugation partner for live cell imaging.

"Turn-on" fluorescent chemosensor for zinc(II) dipodal ratiometric receptor: application in live cell imaging.