Shi J Querard

Le Saux

of intracellular metabolites and proteins in living bacteria

Keywords: fluorogenic probes, fluorescence, protein labeling, biological imaging

Chapter I General Introduction

Bio-imaging, which makes the invisible visible, is the most direct method to reveal complicated and mysterious biological activities in living body. There are two general imaging modalities, clinical modalities, such as magnetic resonance imaging (MRI), X-ray computed tomography (CT), etc. and research modalities, like all kinds of optical microscopes, mass spectrometry imaging, etc., 1 which have significant contribution to the development of basic science in all directions.

Live cell imaging, which unveils cellular events and enables to observe biomolecular processes in real time, has been applied to a wide range of biological researches.

Development of novel reporters and research microscopy techniques has revolutionized the way of visualizing biological activities in cells and accelerate the pace of discovery. Proteins or other biomolecule as the crucial elements of many processes, are normally invisible and usually need to be visualized and tracked in space and time to help biologists to understand their functions. Currently, fluorescence is one of the most widely applied methods of investigation due to its high sensitivity and low experimental cost.

I-1 Fluorescence

Fluorescence is an emission of light from a molecule that has absorbed photons. This phenomenon has been first observed in 1565 by Nicolas Monardes, a Spanish physician and botanist. The term of fluorescence was introduced in 1852 by George Gabriel Stokes. [2][START_REF] Plamont | Small fluorescence-activating and absorption-shifting tag for tunable protein imaging in vivo[END_REF][4]

I-1.1 The principle of fluorescence

The physical principle of fluorescence can be explained by the Perrin-Jablonski diagram (Figure 1.1). The singlet fundamental electronic state is denoted as S 0 , singlet excited electronic states S 1 , S 2 …and the triplet states T 1 , T 2 …Each electronic state contains a range of vibrational and rotational energy levels. After the absorption of photon, electron from S 0 is promoted to the high energy electronic state S 1 . The excited electron is not stable and prone to lose the extra energy to go back to the stable state S 0 through various de-excitation processes, such as internal conversion (IC), intersystem crossing (ISC), fluorescence and phosphorescence. Fluorescence is the emission of photons accompanying the relaxation from S1 to S0.

I-1.2 Absorption, excitation and emission spectra

The absorption of the energy of a photon is an all or none phenomenon. There is no absorption if the photon contains insufficient energy for an electronic transition. On the contrary, absorption occurs when the photon contains enough energy and the excess energy is usually converted into vibrational and rotational energy. Consequently, the absorption of photon by fluorescent molecules can only occur with incident light of specific wavelengths known as absorption bands.

In general, fluorescent molecules undergo electronic transitions with radiation having wavelengths ranging from the ultraviolet to the visible regions of the electromagnetic spectrum. Absorption spectra of fluorescent molecules can be recorded by scanning absorbance A() (a factor that demonstrates the efficiency of absorption) at certain wavelength  with ultraviolet-visible spectroscopy and their absorbance usually follows the Beer-Lambert Law.

Where I 0 () and I() are the intensities of the light received and transited by the absorbing sample; () is the molar absorption coefficient (in L mol -1 cm -1 ), is the absorption path length (in cm) and c is the concentration (in mol L -1 ) of absorbing sample. 2 Excitation spectra of most fluorescent molecules are identical to their absorption spectra.

Generally, the differences of vibrational levels are similar in the ground and excited states, so that the excitation and emission spectra are usually symmetrical. The wavelength of emitted light is longer than the incident light because of the energy loss of vibrational relaxation in the excited state. The difference between the maximum wavelength of excitation and emission spectra is termed as the Stokes shift (Figure 1.2). 2 Excitation and emission spectra can be recorded by scanning fluorescence intensity in a selected spectrum region with a fluorometer at a fixed emission or excitation wavelength. 5 Through recording excitation and emission spectra of fluorescent reporters, one can learn their spectral properties and obtain indispensable information for the spectral region they can be applied in cell imaging. The color of light that we usually speak of can be quantitively described as range of wavelength (Figure 1.3).

I-1.3 Fluorescence quantum yield, lifetime and brightness

The amount of fluorescence emitted by a fluorescent reporter depends on many factors. 2,6 Among them, the fluorescence quantum yield and lifetime are crucial for characterizing fluorescence performance of fluorescent reporter.

Fluorescence quantum yield (Φ) is the ratio of the number of photons emitted to the number of photons absorbed by fluorescent species, which reflects its efficiency of fluorescence emission and is given by:

where k r is the rate constant for radiative deactivation from S 1 to S 0 through fluorescence emission, and k nr is the rate constant for non-radiative deactivation from S 1 to S 0 .

The lifetime of the excited state S 1 is the average time the molecule spends in the S 1 before returning to S 0 and is given by: Usually, one measured the relative quantum yield of fluorescent reporter by comparison with standards whose quantum yields are known, such as fluorescein (0.95±0.03 in 0.1M NaOH), Rhodamine 6G (0.94 in ethanol), etc. 6 The relative quantum yield of fluorescent reporter is calculated by: where the subscript s stands for the standard, A(λ) is the absorbance at the excitation wavelength λ, D is the integrated intensity of emission, and n is the refractive index of the solvent.

The brightness (B) is an important parameter for estimating the fluorescence performance of a fluorescent sample, which is determined by the molar absorption coefficient and the fluorescence quantum yield. Generally, brighter fluorescent reporters increase the signal-tonoise ratio. Moreover, the weaker intensity of light for exciting brighter fluorescent reporters decreases intervention and phototoxicity for hosts. 7 Fluorescence techniques have many impressive advantages compare to other analytical and detective methods, explaining the wide applications of fluorescence in various domains. 2,8 Notably, the development of all kinds of fluorescence microscopies has been boosting

spectacularly the ability of live-cell imaging.

I-1.4 Advantages of fluorescence-based investigative technologies

Fluorescence techniques have many impressive advantages compare to other analytical and detective methods, explaining the wide applications of fluorescence in various domains. 2,8 High sensitivity. Fluorometry is up to 1000 times more sensitive than spectrophotometry. 2 With proper dyes and experimental instruments, the absolute sensitivity of fluorescence detection can reach the limit of single molecules. 6,8 Instead of obtaining average value of ensemble fluorophores in the observing region, behavior and performance of single fluorophore can be recorded by single-molecule fluorescence microscopy, which can provide additional information on the dynamics of fluorescently labeled biomolecule. 6,[START_REF] Moerner | Single-molecule spectroscopy and imaging over the decades[END_REF][START_REF] Kusumi | Tracking single molecules at work in living cells[END_REF] High temporal and spatial resolution. The response of fluorescence techniques could be as fast as 10 -8 -10 -10 s, which is limited by the fluorescence lifetime and the experimental conditions. 8 The spatial resolution is restricted by the diffraction limit of light. Many novel fluorescence super-resolution microscopy technologies have been developed to overcome the light diffraction and to reach nanometric spatial resolution. [START_REF] Hell | Far-field optical nanoscopy[END_REF][START_REF] Betzig | Imaging intracellular fluorescent protein at nanometer resolution[END_REF][START_REF] Gahlmann | Exploring bavterial cell biology with single-molecule tracking and super-resolution imaging[END_REF]

I-2 Fluorescent reporters

Molecules that can generate fluorescence after absorbing photons are known as fluorophores.

Specific fluorophores are needed to track movement of biomolecules for various biological researches. Fluorophores can be endogenous, such as some aromatic amino acids, fluorescent proteins, etc., which naturally exist in organisms and can be studied directly through fluorescence imaging techniques. However, the number of endogenous fluorophores is limited, which restricts application of fluorescence imaging. To conquer this, diversiform exogenous fluorescent probes have been developed and introduced into live cell for biological research.

I-2.1 Organic fluorescent probes

An important landmark in the field of fluorescent probes has been the development of organic fluorescent dyes. They can have some affinity for certain biomolecules or metabolites, and thus visualize locations of biomolecules and reveal biological processes in cells. Innumerous molecular probes with various properties has been synthesized and applied to a large numbers of biological studies. [START_REF] Lavis | Bright building blocks for chemical biology[END_REF][START_REF] Lavis | Bright ideas for chemical biology[END_REF] Two key advantages of the organic fluorescent probes are the small size of these molecules, which will not disturb the natural function of labeled elements, and their marvelous versatility which enables to easily adjust their properties to the need of the experiment through chemical modulations. However, the fact that they show low labeling specificity has encouraged researchers to continue developing novel fluorescent probes for the selective fluorescent labeling of biomolecules, which tremendously boosted imaging techniques based on fluorescence and accelerated the progress of biological research. A substantive leap is the discovery and application of the green fluorescent protein (GFP).

I-2.2 Autofluorescent proteins (AFPs)

GFP is a companion protein of the chemiluminescent protein Aequorin, discovered by Shimomura et al. from the jellyfish Aequorea Victoria. [START_REF] Shimomura | Extraction, purification and properties of aequorin, a bioluminescent protein from the luminous hydromedusan, Aequorea[END_REF] Thirty years later, the first cloning of the gene of GFP [START_REF] Prasher | Primary structure of the Aequorea victoria green-fluorescent protein[END_REF] and the initial experiments of expressing GFP in other organisms [START_REF] Chalfie | Green fluorescent protein as a marker for gene expression[END_REF] opened the new era of bio-imaging. In most applications, AFP works as a fluorescent tag that is genetically fused to a protein of interest (POI), allowing visualizing specifically the fusion protein (Figure 1.4). The high labeling specificity provided by AFPs makes them indispensable in biological research. Since the success of GFP, many GFP-like AFPs with various colors have been developed to cover the spectral needs in multicolor imaging experiments (Figure 1.5). [START_REF] Giepmans | The fluorescent toolbox for assessing protein location and function[END_REF][START_REF] Tsien | The green fluorescent protein[END_REF][START_REF] Chudakov | Fluorescent protein as a toolkit for in vivo imaging[END_REF][START_REF] Shaner | Advances in fluorescent protein technology[END_REF][START_REF] Shaner | Improved monomeric red, orange and yellow fluorescent proteins derived from Discosoma sp. red fluorescent protein[END_REF][START_REF] Wang | Evolution of new nonantibody proteins via iterative somatic hypermutation[END_REF][START_REF] Dean | Advances in fluorescence labeling strategies for dynamic cellular imaging[END_REF]

I-2.2.1 Applications of AFPs

Labeling

The first type of application is labeling. Genetic manipulations allow to fuse AFP to a protein of interest or a binding domain for a given biomolecule, which enables to obtain information about location, movement, function of biomolecules and can provide us with information on biological constituents (organelle, subcellular structure, cell, organism, tissue, etc.).

Apart from conventional labeling, AFPs have been used to develop novel techniques for biological detection and analysis.

Photobleaching techniques

Photobleaching is a common and undesirable phenomenon during the observation of fluorophore-labeled biomolecule through light irradiation. However, applying photobleaching as a disturbance factor can provide information about the mobility of biomolecules, which is important to understand their activities and functions in cell. 7,[START_REF] Lippincott-Schwartz | Photobleaching and photoactivation: following protein dynamics in living cells[END_REF][START_REF] Day | The fluorescent protein palette: tools for cellular imaging[END_REF] The most obvious strategy is fluorescence recovery after photobleaching (FRAP

Förster resonance energy transfer (FRET)

FRET is an important technique to study protein-protein interactions. Two AFPs are chosen AFPs fluoresce only after the formation of the fluorophore. The multi-step maturation (Figure 1.8) usually requires the presence of oxygen and takes tens of minutes or hours, [START_REF] Cubitt | Understanding, improving and using green fluorescent proteins[END_REF][START_REF] Heim | Wavelength mutations and posttranslational autoxidation of green fluorescent protein[END_REF] which prevents their applications in anoxic conditions or for studying fast biological processes.

Although AFPs with improved brightness, [START_REF] Heim | Improved green fluorescence[END_REF] better photostability [START_REF] Shaner | Improving the photostability of bright monomeric orange and red fluorescent proteins[END_REF] or faster maturation [START_REF] Evdokimov | Structural basis for the fast maturation of Arthropoda green fluorescent protein[END_REF] have been developed, we still expect further improvement that could combine the high specificity of AFP and versatility of small fluorescent molecules in the same fluorescent probe system.

The most impressive developments are the site-specific labeling techniques, in which peptidic tags are used as genetic anchors reacting specifically with synthetic probes.

I-2.3 Site-specific labeling techniques I-2.3.1 Biarsenical-tetracysteine tagging system

The first tag developed for site-specific protein labeling was the tetracysteine motif, a short peptide usually composed of six amino acids CCPGCC that can bind with biarsenical dyes. [START_REF] Adams | New biarsenical ligands and tetracysteine motifs for protein labeling in vitro and in vivo: synthesis and biological applications[END_REF] The principle of the binding is the well-known interaction of arsenic with thiol groups. The first developed biarsenical dye, FlAsH, is a fluorescein derivative with two As(III) substituents, which is weakly fluorescent as an adduct with two 1,2-ethanedithiol (EDT) but becomes green fluorescent after binding with the tetracysteine tag (Figure 1.9). [START_REF] Griffin | Specific covalent labeling of recombinant protein molecules inside live cells[END_REF][START_REF] Griffin | Fluorescent labeling of recombinant proteins in living cells with FlAsH[END_REF] Analogs have been developed for expending the spectral properties. [START_REF] Adams | New biarsenical ligands and tetracysteine motifs for protein labeling in vitro and in vivo: synthesis and biological applications[END_REF][START_REF] Gaietta | Multicolor and electron microscopic imaging of connexin trafficking[END_REF] ReAsH (Figure 1.9) as the most useful one is a derivative of the red fluorophore resorufin and could be applied in pulsechase imaging with FlAsH. [START_REF] Gaietta | Multicolor and electron microscopic imaging of connexin trafficking[END_REF] Although the small size of the peptide tag and the various biarsenical dyes makes it one of the best established site-specific labeling systems, it still needs to be improved in many aspects like the signal-to-background ratio, the binding specificity, the need of antidotes to reduce the toxicity and the requirement of a reducing environment. 36 Since the development of biarsenical-tetracysteine tagging system, there have been many

efforts to create new site-specific labeling systems with better properties. SNAP-tag, CLIP-tag and Halo-tag are three outstanding representatives which are all commercially available now.

I-2.3.2 Self-labeling tags: SNAP-tag, CLIP-tag, Halo-tag

SNAP-tag is a mutant of the human DNA repair protein O 6 -alkylguanine-DNA alkyltransferase (AGT) developed by the group of Johnsson. 40 -43 It can transfer the functionalized benzyl group of the O 6 -benzylguanine (BG) derivative to its active site cysteine and thus achieves covalent and irreversible labeling of fusion protein (Figure 1.10).

The versatility of the developed BG derivatives enables to easily adapt SNAP-tag functionalities to different conditions for visualizing proteins in vitro and in vivo. [START_REF] Kindermann | Synthesis and characterization of bifunctional probes for the specific labeling of fusion proteins[END_REF][START_REF] Keppler | Labeling of fusion proteins of O 6 -alkylguanine-DNA alkyltransferase with small molecules in vivo and in vitro[END_REF][START_REF] Gronemeyer | Adding value to fusion proteins through covalent labelling[END_REF] To meet the need for multiplexed imaging, the group of Johnsson developed CLIP-tag [START_REF] Gautier | An engineered protein tag for multiprotein labeling in living cells[END_REF] , which shows substrate specificity orthogonal to SNAP-tag. CLIP-tag is an engineered mutant of AGT that works as SNAP-tag but reacts specifically with O 2 -benzylcytosine (BC) derivatives (Figure 1.10). These two tags have been used to label two different proteins in living cells for simultaneous pulse-chase experiments [START_REF] Gautier | An engineered protein tag for multiprotein labeling in living cells[END_REF] and also used to cross-link interacting proteins to detect and characterize protein-protein interactions [START_REF] Gautier | Selective cross-linking of interacting proteins using self-labeling tags[END_REF] .

The Halo-tag is engineered from a bacterial halogen dehydrogenase. The carbon halogen bond of the functionalized chloroalkane derivative could be cleft by the Halo-tag and thus the fluorescent ligand covalently binds with the Halo-tag fusion protein (Figure 1.10). 49 The labeling specificity and the easy chemical modification of the substrates make these selflabeling systems very attractive. However the imaging of all these systems can show fluorescence background because of the excess of substrates and nonspecific binding.

Therefore extensive washing steps are requested before imaging, which may limit the temporal resolution of the experiments.

I-2.3.3 Fluorogenic site-specific labeling systems

To improve the imaging contrast, there have been some developments of fluorogenic substrates for these self-labeling systems. Fluorogenic chromophores (so-called fluorogens) . [START_REF] Klymchenko | Solvatochromic and fluorogenic dyes as environment-sensitive probes: design and biological applications[END_REF] These SiR probes enable imaging fusion proteins with high sensitivity at 640-650 nm, a wavelength range where cellular autofluorescence is weak and phototoxicity is minimal. 53 These fluorogenic site-specific labeling systems eliminate the high fluorescence background, the common disadvantage for chemical labeling and save the time for washing steps.

However, the relatively large sizes of SNAP-tag (20kDa), CLIP-tag (20kDa) and Halo-tag (34kDa) and the long time for complete labeling still may hinder their applications in some cases. Actually, there have been many efforts to combine different fluorogens with different small genetically encoded tags to create novel fluorogen based specific labeling systems with high temporal resolution.

I-2.4 Fluorogen-activating proteins (FAPs)

Fluorogen-activating proteins (FAPs) engineered from libraries of human single-chain antibodies (scFvs) bind non-covalently fluorogenic derivatives of malachite green (MG) and thiazole orange (TO) with nanomolar affinity and generate huge enhancement of their red and green fluorescence, respectively. [START_REF] Szent-Gyorgyi | Fluorogen-activating single-chain antibodies for imaging cell surface proteins[END_REF] MG and TO are well-known molecular rotors that are poorly fluorescent in solution because of internal rotation; however in constrained environments, the internal rotations are slowed down, which strongly enhance their fluorescence intensities. [START_REF] Klymchenko | Solvatochromic and fluorogenic dyes as environment-sensitive probes: design and biological applications[END_REF] FAPs were shown to be suitable to label proteins without the need of wash-out process. Cell impermeant MG-2p and TO1-2p enable labeling of only the FAPtagged proteins at the plasma membrane, while the cell permeant fluorogen MG-ester allows lighting up proteins in the secretory pathway within several minutes after addition (Figure 1.12). 55 Various fluorogens have been developed to bind the single scFv protein K7 which shows huge versatility and accept different fluorogenic dyes. [START_REF] Ozhalici-Unal | A rainbow of fluoromodules: A promiscuous scFv protein binds to and activates a diverse set of fluorogenic cyanine dyes[END_REF][START_REF] Shank | Enhanced photostability of genetically encodable fluoromodules based on fluorogenic cyanine dyes and a promiscuous protein partner[END_REF] Dimethylindole Red (DIR) and its analogues (unsymmetrical cyanines) were developed to bind the FAP K7 and expand the spectral properties of FAPs by giving complexes with emission wavelengths ranging from the blue (450 nm) to near-IR (750 nm) (Figure 1.13). [START_REF] Ozhalici-Unal | A rainbow of fluoromodules: A promiscuous scFv protein binds to and activates a diverse set of fluorogenic cyanine dyes[END_REF] These dyes were further modified (the -CN-DIR and the fluorinated TO (Figure 1.13)) to improve the photostability of the FAP system. [START_REF] Shank | Enhanced photostability of genetically encodable fluoromodules based on fluorogenic cyanine dyes and a promiscuous protein partner[END_REF] The following development of blue FAP systems based on blue fluorogenic cyanine dyes (OTB-SO 3 ) (Figure 1.13) 58 and the far-red FAP systems based on the ester of malachite green dyes (MGe) (Figure 1.13) 59 makes emission colors of FAPs cover almost the whole visible spectrum, which is versatile for multiplexed imaging. However, the fact that single-chain antibodies contain internal disulfide bonds that form only in non-reducing environment limits their applications to non-reducing environments. [START_REF] Szent-Gyorgyi | Fluorogen-activating single-chain antibodies for imaging cell surface proteins[END_REF] To overcome this limitation, disulfide-free variants FAPs has been developed, which enables the labeling of FAP systems in various environment in live cells. 59

I-2.5 PYP-tag

The To further improve the properties of PYP-tag labeling system, they created mutants of the PYP-tag and chemically modified fluorogens by considering the electrostatic interactions of the protein tag with the fluorogen and the pK A of the leaving group. 63,64 Fluorogens that could generate different colors have also been developed for multicolor imaging. 65 The small size of the PYP-tag and the quick wash-free imaging ability make PYP-tag very attractive for fluorescence imaging in living cells.

I-2.6 Cellular retinoic acid binding protein II (CRABPII)

The cellular retinoic acid binding protein II (CRABPII) is a binding and transport protein that binds cellular retinoic acids and thus transport them inside the cell cytoplasm. [67][68][START_REF] Chaudhuri | Structures of cellular retinoic acid binding proteins I and II in complex with synthetic retinoids[END_REF] Its small size (15.6 kDa), relatively large binding pocket that could binds with various synthetic retinoids (Figure 1.17) and tolerance of mutations makes it a promising protein scaffold for the engineering of fluorogenic fluorescent protein tag. [START_REF] Yapici | Turn-on' protein fluorescence: in situ formation of cyanine dyes[END_REF] The group of Borhan has developed a novel fluorogenic protein labeling system by reengineering CRABPII. A non-fluorescent precursor merocyanine aldehyde reacts with the active site Lys residue of CRABPII-tag to generate the fluorescent protonated Schiff base which also displays a huge red-shift of absorption and a significant increase of extinction coefficient compare to the precursor merocyanine aldehyde (Figure 1.18). The chromophore is planar and restricted in the protein cavity, which enhance its fluorescence. [START_REF] Yapici | Turn-on' protein fluorescence: in situ formation of cyanine dyes[END_REF] .

I-2.7 Spinach

Except for visualizing proteins, fluorogenic labeling has also been applied to other genetically encoded biomolecules, such as RNA. There have been many developments of fluorogenactivating RNA aptamer. [START_REF] Babendure | Aptamers switch on fluorescence of triphenylmethane dyes[END_REF][START_REF] Sparano | A strategy for the Development of small-molecule-based sensors that strongly fluoresce when bound to a specific RNA[END_REF][START_REF] Sando | Transcription monitoring using fused RNA with a dye-binding light-up aptamer as a tag: a blue fluorescent RNA[END_REF][START_REF] Constantin | Synthesis of new fluorogenic cyanine dyes and incorporation into RNA fluoromodules[END_REF][START_REF] Armitage | Imaging of RNA in live cells[END_REF][START_REF] Rath | Genetically encoded tools for RNA imaging in living cells[END_REF] However the first fluorogen-based RNA aptamer that could efficiently image RNA in living cells is the Spinach system (Figure 1.19) which has been developed by the group of Jaffrey. [START_REF] Paige | RNA mimics of green fluorescent protein[END_REF] Spinach relies on the binding and activation of analogs of HBI, the chromophore of GFP. As presented before, GFP becomes fluorescent only after the formation of HBI which is a well-known fluorogenic chromophore. HBI is only fluorescent when encased in GFP. Denaturation of GFP leads to a loss of fluorescence. [START_REF] Paige | RNA mimics of green fluorescent protein[END_REF] Inspired by the GFP, the group of Jaffrey designed and synthesized several fluorogenic analogs of HBI and selected RNA aptamers that could bind specifically with the fluorogen from a library of random RNA molecules. [START_REF] Paige | RNA mimics of green fluorescent protein[END_REF] To improve the properties of Spinach, they developed Spinach2, which shows enhanced folding and thermal stability [START_REF] Strack | A superfolding Spinach2 reveals the dynamic nature of trinucleotide repeat-containing RNA[END_REF] and they selected Broccoli through a fluorescence activated cell sorting (FACS) based directed evolution approach (Figure 1.20) [START_REF] Filonov | Rapid selection of an RNA mimic of green fluorescent protein by fluorescence-based selection and directed evolution[END_REF] . The development of HMBR was inspired by GFP. HMBR is an analog of HBI and is based on a push-pull structure. The electron-donating phenol group interacts with electron-withdrawing rhodanine group through a conjugated system. HMBR (pK a =8.4) stays almost all in protonated form at pH 7.4 showing no fluorescence. However, HMBR binds with FAST in its deprotonated form. The binding induced deprotonation of the phenol group generates phenolate with higher electron-donating effect, which explains the absorption red shift.

Meanwhile, constrained environment in the protein cavity restrains the conformational freedom of HMBR and thus activates its fluorescence.

FAST is an engineered variant of PYP. The chromophore of PYP, parahydroxycinnamoyl Compared to other fluorogen-based labeling systems, FAST has many attractive advantages:

The protein tag is monomeric and small (14 kDa), which minimize its hindrance for normal activities of fusion protein; FAST works as soon as it is folded and the labeling of FAST fusion protein occurs almost immediately after addition of fluorogen, which enables its application for real-time imaging; The binding between FAST and its fluorogen is reversible and shows rapid exchange dynamics, which makes labeling of FAST highly tunable. It has been proved that FAST can be rapidly switched on and off by addition and removal of HMBR and this process is repeatable. The controllable labeling opens new opportunities for multiplexed imaging. These advantages make FAST a very promising protein tag. My PhD aimed at extending the possibilities offered by FAST through the engineering of new fluorogens and additional protein engineering. The first step was the development of novel fluorogens. HMBR is a push-pull system. The electron-donating phenol group interacts with the electron-withdrawing rhodanine group through a conjugated system. Molecules with such push-pull structure are known as environment-sensitive dyes [START_REF] Klymchenko | Solvatochromic and fluorogenic dyes as environment-sensitive probes: design and biological applications[END_REF] . Spectral properties of these molecules are determined by the electron-donating or electron-withdrawing abilities of their electron donors or acceptors and the length of the electronic conjugation. On the basis of this push-pull structure, we engineered HMBR analogues by modifying either their electrondonating or electron-withdrawing groups or the length of the conjugation paths (Figure1.23).

I-3 Objectives

Modification of the electron-donating group allowed us to generate fluorogens binding FAST with red-shifted emission (Chapter II). Increase of the conjugation and variation of the electron-withdrawing group enabled us to generate fluorogens with far-red emission when bound to engineered variants of FAST (Chapter III). Finally, introduction of charged groups on the heterocycle of HMBR allowed us to render HMBR membrane-impermeant for the selective labeling of membrane proteins (Chapter IV).

Based on this chapter, we have written a relative review.

Fluorogenic labeling strategies for biological imaging.

International Journal of Molecular Sciences

Chapter II Expansion of the spectral properties of FAST

This chapter is based on the following article: We designed and synthesized a collection of HMBR analogs with various aromatic substituents in different positions of the phenol ring (Figure 2.1). New fluorogens were obtained by condensation of the starting aldehydes and rhodanine in water or refluxing ethanol in presence of piperidine. Their abilities to bind FAST were verified by characterizing their spectral properties alone or in the presence of FAST. We also tested their binding affinity by measuring thermodynamic dissociation constants (K D ). As expected, we observed fluorescence activation and absorption red shift in the presence of FAST, which demonstrated that they all kept the main features of the FAST approach. These FAST:fluorogen complexes displayed various spectroscopic behaviors. Compared to HMBR, HBR-3E in which an ethyl group replaces the methyl group of HMBR binds to FAST with higher affinity and forms complex displaying almost identical absorption and emission properties but reduced fluorescence brightness (See Article 1-Table 1). As brightness is directly related to the immobilization of the fluorogen within the protein cavity, we next examined the effect of an additional methyl group either in ortho-position or metaposition of the phenolic hydroxyl group. HBR-2,5DM and HBR-3,5DM bind tightly to FAST and form complexes displaying a 15-20 nm red shift in absorption and a 10-20 nm red shift in emission with improved fluorescence brightness compared to FAST:HMBR (See Article 1-Table 1). Of particular interest, FAST:HBR-3,5DM is more than two times brighter than FAST:HMBR, reaching fluorescence efficiency of enhanced green fluorescent protein (EGFP).

To further shift absorption and emission to the red, we next replaced the methyl group of HMBR with alkoxy groups (methoxy or ethoxy group), which are common electron donors.

HBR-3OM and HBR-3OE bind to FAST with affinities comparable to HMBR and form complexes displaying a 15 nm red shift in absorption and a 20 nm red-shift in emission. While FAST:HBR-3OE was less bright than FAST:HMBR, FAST:HBR-3OM exhibited greater brightness (See Article 1-Table 1). We next tested the introduction of a second methoxy group in ortho-position of the phenolic hydroxyl group. The resulting complex, FAST:HBR-3,5DOM, displayed a 40-nm red shift in absorption and a 60-nm red shift emission, and improved fluorescence brightness compared to FAST:HMBR (See Article 1-Table 1). These spectral modifications shift the absorption to the green and the emission to the orange-red, providing thus a new color for fluorescence imaging. In terms of fluorescence performance, FAST:HBR-3,5DOM was as bright as the red fluorescent protein mCherry.

In sum, we obtained four new fluorogens (HBR-3OM, HBR-2,5DM, HBR-3,5DM and HBR-3,5DOM) enabling to tune the fluorescence color of FAST from green-yellow to orange and red with higher brightness. Their pK a are between 8.3-8.7 (See Article 1-Table S1)

ensuring that they stay almost fully protonated at pH 7.4 with no fluorescence when free in solution. Moreover the range of K D (~0.1-1M) (See Article 1-Table 1) keeps their capabilities of reversible labeling with FAST-tagged proteins in living cells, while ensuring selective labeling.

We next verified their performance for labeling protein in living cells. Cytotoxicity testing proved that they were non-toxic for mammalian cells (See Article 1-Figure S2). Then we treated HeLa cells expressing FAST in different cell localizations (cytoplasm, nucleus or mitochondria) with the new fluorogens (HBR-3OM, HBR-3,5DM or HBR-3,5DOM). We observed specific fluorescence in the expected localizations through confocal microscopy (See Article 1-Figure 2). Their abilities of reversible labeling were examined by labeling HeLa cells expressing nuclear FAST repeatedly with HBR-3OM, HBR-3,5DM or HBR-3,5DOM solutions and then reversing the labeling by removing the fluorogencontaining solutions followed by a wash with pH 7.4 phosphate buffer saline (PBS) (See and calculating the degree of anti-correlation of the green and red fluorescence signals, the correct localization of FAST was obtained (Article 1-Figure 4b). This experiments showed that dynamic multi-color protein labeling strategy could allow overcoming the limitation of observation channels, opening new opportunities for multiplexed imaging.

The labeling of FAST-tagged proteins with HMBR or HBR-3,5DOM is not orthogonal because they share the same protein tag. Ongoing projects in our team aim at selecting FAST variants that display orthogonal selectivity and eventually better spectral properties. Directed evolution approaches based on yeast display technology and Fluorescence-Activated Cell Sorting (FACS) are used to isolate variants from yeast-displayed FAST variant libraries (See also Chapter III). We are expecting FAST variants that could bind specifically and orthogonally with HMBR and HBR-3,5DOM with an improved brightness.

To further expand the spectral properties of FAST to the far-red edge of the visible spectrum, we also engineered new fluorogens and protein tags, which will be discussed in the Chapter III.

III-1 Molecular engineering of red emitting fluorogens

This section is based on the following article:

Design and characterization of red fluorogenic push-pull chromophores holding great potential for biosensing and imaging.

Organic and Biomolecular

III-1.1 Presentation of article 2

The fluorogens developed for FAST are based on a push-pull conjugated structure, which consist in an aromatic electron-donating part and a heterocyclic electron-withdrawing terminal group interacting through a conjugated system. For push-pull fluorophores, effects of electron-donating and electron-withdrawing character and the extension of the conjugated system influence their spectral properties. We designed two series of putative fluorogens by combining two electron donors, the 4-hydroxyphenyl (series 1) or the 4-dimethylamino-phenyl group (series 2) with various heterocyclic electron acceptors through longer conjugated system (with an additional double bond compared to previous fluorogens) (Figure 3.1) for expending furthest their spectral properties to red. Apart from engineering red fluorogens for FAST approach, we have demonstrated in this article the general idea and method to develop and characterize potential red fluorogens in this article. New fluorogens were synthesized by condensation of the starting aldehydes and heterocycles in refluxing ethanol in presence of amine base (See Article 2-Figure 2). For series 2, the starting aldehyde is commercially available while the starting aldehyde for series 1 was obtained from a methoxymethyl (MOM)-protected 4-hydroxybenzaldehyde which underwent a Wittig oxyprenylation followed by acid hydrolysis (See Article 2-Figure 2). Next, we characterized their spectral properties (absorption and fluorescence emission) in aqueous buffer and in different conditions mimicking environment changes in protein cavity to see if they possess potential fluorogenic properties.

Chromophores of series 1 have two forms, protonated (phenol group) and deprotonated (phenolate group) form. Their pK a are between 8.8-8.9 (See Article 2-Table 1 and Figure 3) ensuring that they stay almost fully protonated at pH 7.4. The phenolate displays higher electron-donating effect than a phenol group leading to a red-shift of the absorption. These acido-basic properties can be used to increase the fluorogenic response, since binding to a complementary cavity stabilizing the negatively charged phenolate could result in a large red-shift in absorption at physiological pH, as previously demonstrated in the case of FAST and the HBR family 1 (See also Chapters I and II). We tested their absorption and emission properties at pH 6 (fully protonated) and pH 10 (fully deprotonated) to see the differences between these two forms. Deprotonation of phenol group induced 20-65 nm red shift in absorption and 55-95 nm red shift in emission, shifting the absorption from ultra violet or violet to violet or blue and the emission from yellow or orange to orange or red (See Article 2-Table 1 and Figure 4). The two forms displayed weak fluorescence (fluorescence quantum yield < 2%, see Article 2-Table 1). To test their fluorogenic response upon conformational locking, we used aqueous buffer at pH 10 complemented with 40% glycerol. Glycerol increases solution viscosity and thus restricts the motion of chromophores, mimicking environment changes in protein cavities. We observed about two-fold increase of fluorescence quantum yield (See Article 2-Table 1), which proved their capability of fluorescence enhancement upon conformational restriction. From our previous work 1 , this behavior allowed us to anticipate larger fluorogenic responses upon binding within a tailored protein cavity. Chromophores of series 2 absorbed violet or blue light and fluoresced weakly in red (fluorescence quantum yield < 1%) in aqueous buffer at pH 7.4 (See Article 2-Table 2 and Figure 5). In viscous buffer (in presence of 40% glycerol), they also displayed about two-fold increase of fluorescence quantum yield (See Article 2-Table 2). Moreover, solvato(fluo)chromic properties tests revealed their negative solvatochromism properties (See Article 2-Figure 6), which could also be used to obtain absorption red shift upon entering lowpolarity protein cavities. In summary, the two series of chromophores are all potential red fluorogens for FAST approach because they display fluorescence activation upon motion restriction and potential of red shift in absorption through binding-induced ionization of auxochromic group or negative solvatochromism. In particular, these fluorogens exhibit high Stoke shifts from 150 nm to 260 nm, opening great perspectives for multiplexed imaging with a single excitation source.

Introduction

We have presented above the general method to develop red-emitting fluorogens and obtained several potential red-emitting fluorogens for fluorogenic labeling. In this part, the general idea for engineering protein tags able to bind and activate some of these new fluorogens, will be introduced.

Protein engineering is commonly conducted by two strategies, rational protein design and directed evolution. Rational protein design aims at generating desired variants through sitedirected mutagenesis of original protein gene. This strategy requires full knowledge of the structure and function of the original protein, which is important to design mutants with expected properties. However, influence of structural modifications on functional changes is difficult to predict. Another strategy is directed evolution for which desired protein variant will be selected from large protein libraries generated by mutagenesis of the original protein gene. Compared to rational design, directed evolution does not need to understand the original protein in detail and the large numbers of mutants increase the chance to obtain protein variants with desired properties. However, screening out rare desired mutants from mutant libraries takes a lot of time and effort. High throughput selection methods are thus required for accomplishing directed evolution efficiently. Before starting to screen complementary protein tags for putative fluorogens, we needed to identify protein scaffolds containing cavity that could possibly accept the fluorogens and activate their fluorescence. PYP had originally been chosen as protein scaffold to develop FAST because its chromophore parahydroxycinnamoyl (HC) shared structure similarity with HMBR. In Section III-1, the red-emitting fluorogens we developed were analogues of HMBR with an additional double bond. So firstly we tested their binding abilities with FAST. Except for 1c (4-hydroxyphenyl)allylidene-rhodanine (HPAR), which showed a little fluorescence activation by interacting with FAST, the other fluorogens did not exhibit any binding affinity with FAST. We thus decided to focus first on the evolution of FAST variants able to bind the fluorogenic HPAR and to activate its red fluorescence. We built a library of FAST variants by random mutagenesis using error-prone PCR. Error-prone PCR is a common technique for random mutagenesis, which generates mutations during PCR process by using error prone DNA polymerases. We were able to identify from this libraries FAST variants able to bind and activate the red fluorescence of HPAR.

III-2.2 Results and discussions

Construction of the library of FAST variants.

Our library of FAST variants was generated by random mutagenesis of FAST gene by errorprone PCR. A mutation rate of 3.8 nucleotides/gene was obtained after mixing PCR reactions with different error rates varied by adjusting the amount of template gene. The PCR product was cloned into the pCTCON2 plasmid from Wittrup's lab, allowing the expression of the FAST variants in fusion to Aga2p. The resulting plasmids were transformed into DH10B E.

coli cells with a yield of 10 6 transformants. DNA was extracted and then transformed into EBY100 yeast strain using a large-scale high-efficiency transformation protocol 5 , leading to 2 × 10 6 transformants. In sum, we constructed a FAST variant library containing about 10 6

FAST variants.

Choice of HPAR for screening protein tag.

HPAR was considered to be the most promising for engineering red-emitting FASTs. Its physicochemical properties in aqueous buffer and conditions mimicking environmental changes in protein cavity showed that it could be possible to preserve fluorescence activation and absorption red shift by binding with complementary protein tag (See Section III-1).

Moreover, its structure shares the most similarity with that of HMBR, allowing us to envision evolving the binding properties of FAST. We showed that it was the only one among the different tested fluorogens that exhibited a binding ability albeit weak with FAST. In aqueous buffer, its deprotonated form fluoresces weakly red light (maximal emission wavelength:

655nm), demonstrating great potential for the generation of red-emitting FASTs. It was furthermore shown to be non-toxic for mammalian cells even at high concentration (See Article 2-Table 3) and not to undergo any non-specific fluorescence activation in living cells (See Article 2-Figure 7). For all these reasons, we chose HPAR as fluorogen candidate to engineer new protein tags for the development of red-emitting FASTs.

Screening of new protein tags

We screened the yeast surface-displayed FAST library in the presence of 10 M HPAR by high-throughput FACS. The level of protein expression on the yeast surface was normalized by immunolabeling the expressed proteins with an Alexa647-conjugated antibody before cell sorting. According to the spectral properties of HPAR in different conditions, the prospective fluorogen:protein tag complexes were expected to absorb blue-green light and fluoresce red.

We thus excited yeast cells incubated in HPAR solution with 488 nm laser and detected and

Physicochemical characterization

The selected variants were expressed in Escherichia coli and purified by affinity chromatography. So far, we tested the spectral properties of two protein variants:HPAR complexes, with clone 1and clone 3. Unlike FAST which showed little binding affinity with HPAR, these protein variants could bind HPAR more tightly and formed red-emitting complexes preserving the features of fluorescence activation and absorption red shift upon binding (Figure 3.5, Table 3.1). Compared to FAST:HBR, protein variants:HPAR complexes displayed 34-42nm red shift in absorption and a 108nm red shift in emission (Figure 3.6, Table 3.2), which verified the bathochromic effect induced by the elongation of the conjugated system. Moreover, the new complexes displayed 9-17 nm blue shift in absorption and 34-36nm red shift in emission compared to the previous red-emitting FAST:HBR-3,5DOM, generating a real red-emitting ( em ~635 nm) fluorescent complexes with a huge Stokes shift (~130nm) (Figure 3.6, Table 3.2) which is attractive for multicolor imaging with a single excitation source.

III-2.3 Conclusion and perspective

We have selected 4 FAST variants activating fluorescence of HPAR from the FAST variant library. Two protein variants were purified and spectral properties of variant:HPAR complexes were characterized. Actually, redder fluorescent complexes which displayed 35nm red shift in emission compared to the previous red-emitting FAST:HBR-3,5DOM were obtained with a higher Stokes shift (~130nm), which showed great potential for multicolor imaging with a single excitation source.

The spectral differences between FAST:HBR and new red-emitting complexes proved that the strategy of elongating the conjugated system of HBR worked for extending spectral properties to the red. We envision to engineer HPAR analogues to further shift their spectral properties to red and improve the binding affinity and brightness of these new red-emitting complexes.

The other two variants will later be expressed and then spectral properties of FAST variant:HPAR complexes will be characterized. Variants which are able to activate specifically the red fluorescence of HPAR preserving unique feature of FAST approach will be identified and compared with the previous characterized two complexes. Next the performance of the best red-emitting complex will be tested in living cells.

III-2.4 Materials and Methods

Yeast display library construction

The gene of FAST was randomly mutagenized by error-prone PCR using the GeneMorph II Random Mutagenesis Kit (Agilent). The error rate of the PCR was varied by using either 1 or 10 ng template gene. The mixed PCR product was digested with Nhe I and BamH I, and then ligated in pCTCON2 using Nhe I / BamH I restriction sites. Large-scale transformation into DH10B E. coli cells was performed by electroporation. The DNA was then maxiprepped and retransformed into EBY100 yeast strain using a large-scale high-efficiency transformation protocol 5 .

Yeast cell sorting

Yeast library (about 1× 10 The sorted cells were collected in SD, grown overnight (30°C, 240 rpm) and spread on SD plates (SD supplemented with 182 g/L sorbitol, 15 g/L agar). Plates were incubated for 60 h at 30°C. The cell lawn was collected in SD supplemented with 30% glycerol, aliquoted and frozen or directly used in the next round.

Molecular Biology

The plasmids pAG291 and pAG293 driving bacterial and in vitro expression of clone 1and clone 3, respectively, with an N-terminal His-tag under the control of a T7 promoter were obtained by inserting the gene encoding clone 1 or clone 3 between Nhe I and Xho I restriction sites in the pET28a vector.

Bacterial expression and protein purification Expression vectors were transformed in

Rosetta(DE3)pLysS E. coli (New England Biolabs). Cells were grown at 37°C in Lysogeny Broth (LB) medium complemented with 50 μg/ml kanamycin and 34 μg/ml chloramphenicol to OD 600nm 0.6. Expression was induced for 4 h by adding isopropyl β-D-1thiogalactopyranoside (IPTG) to a final concentration of 1 mM. Cells were harvested by centrifugation (4,300 × g for 20 min at 4°C) and frozen. The cell pellet was resuspended in lysis buffer (phosphate buffer 50 mM, NaCl 150 mM, MgCl 2 2.5 mM, protease inhibitor, DNase, pH 7.4) and sonicated (5 min at 20 % of amplitude). The lysate was incubated for 2 h at 4 °C to allow DNA digestion by DNase. Cellular fragments were removed by centrifugation (9,300 × g for 1h at 4°C). The supernatant was incubated overnight at 4°C under gentle agitation with Ni-NTA agarose beads in phosphate buffered saline (PBS) (sodium phosphate 50 mM, NaCl 150 mM, pH 7.4) complemented with 10 mM Imidazole.

Beads were washed with 20 volumes of PBS containing 20 mM Imidazole, and with 5 volumes of PBS complemented with 40 mM Imidazole. His-tagged proteins were eluted with 5 volumes of PBS complemented with 0.5 M Imidazole, followed by dialysis with PBS.

Physical Chemistry experiments pH measurements were performed on a standard pH meter PHM210 Radiometer Analytical (calibrated with aqueous buffers at pH 4 and 7 or 10) with a Crison 5208 Electrode (Barcelona, Spain). UV/Vis absorption spectra were recorded in 1 cm × 1 cm quartz cuvettes (Hellma) on a diode array UV/Vis spectrophotometer (Evolution array, Thermo Scientific). Corrected fluorescence spectra upon one-photon excitation were recorded with a Photon Technology International QuantaMaster QM-1 spectrofluorimeter (PTI, Monmouth Junction, NJ) equipped with a Peltier cell holder (TLC50, Quantum Northwest, Shoreline, WA). The overall emission quantum yields after one-photon excitation  were determined as previously described 1 . Previous fluorogens for FAST are all highly membrane-permeant. In order to extend the properties of FAST and provide investigators with the possibility of selectively detecting FAST-tagged cell-surface proteins, we sought for novel fluorogens able to form fluorescent complexes with FAST, but incapable of crossing the plasma membrane. To reduce the ability to penetrate the plasma membrane and improve cell exclusion, we considered functionalizing previous HMBR analogues with a carboxylate, a negatively charged group well known to reduce plasma membrane penetration. We have presented in Chapter II the general method to tune spectral properties of FAST approaches by engineering HMBR analogues with various substituents on their phenol rings. It was shown that aromatic substituents are one of determinants for binding with FAST and for spectral properties of formed FAST:fluorogen complexes, as they can play a major role in immobilizing the fluorogen within FAST cavity.

III-2.5 Reference

Here we tried to develop cell impermeant fluorogens by modifying the rhodanine group of HMBR. An acetate acid was added on the NH position of the rhodanine, expected to create cell impermeant fluorogen and preserve its binding ability with FAST. At pH 7.4, the acetate acid will be deprotonated, staying as acetate with a negative charge. Moreover it has been shown that HBRAA (Figure 4.1) displayed binding affinity with FAST in the initial research for brighter FAST fluorogens [START_REF] Plamont | Small fluorescence-activating and absorption-shifting tag for tunable protein imaging in vivo[END_REF] . The results has also shown that FAST:HBRAA complex was not as bright as desired. In this chapter, we verified that the strategy of adding negative charged acetate on the rhodanine group was feasible to generate cell impermeant fluorogen for FAST and also tried to improve the brightness of cell impermeant FASTs by fluorogen engineering through the same strategy presented in Chapter II. We present in particular HBRAA-3E exhibiting good binding affinity with FAST and forming fluorescent complex with brightness acceptable for cell imaging. This new membrane-impermeant fluoorgen has been applied to selectively image surface-exposed FAST-tagged proteins. It opens great prospects to efficiently obtain spatial information on FAST-tagged proteins by using spectrofluorimetry, which is promising for efficiently screening drugs to treat trafficking related diseases.

IV-2 Results and discussion

Fluorogen design

HMBR is mainly neutral at physiological pH (the phenol pK A is 8.7) and highly membranepermeant [START_REF] Plamont | Small fluorescence-activating and absorption-shifting tag for tunable protein imaging in vivo[END_REF] . To reduce its ability to penetrate the plasma membrane and improve its cell exclusion, we functionized its rhodanine head with an acetate acid group which would be deprotonated and stayed as negative charged acetate at pH 7.4. Commercially available rhodanine acetic acid was condensed with 4-hydroxy-methylbenzaldehyde to provide HBRAA-3M (Figure 4.2), an HMBR analog bearing an additional acetate acid group on the rhodanine head. The resulting FAST:HBRAA-3M complex had a similar brightness (Table 4.1) as the FAST:HBR complex (the first FAST:fluorogen complex before the development of FAST:HMBR, brightness 4000). A simple cell permeability test has been done by imaging yeast cells expressing Aga2p-FAST at the surface and HeLa cells expressing FAST-mCherry in the cytoplasm incubated with HBR or HBRAA-3M (Figure 4.

3). HBR labeled not only

Aga2p-FAST at the yeast cell surface but also Aga2p-FAST in the process of transportation to cell surface, while HBRAA-3M seemed only labeled Aga2p-FAST at the yeast cell surface (Figure 4.3). For HeLa cells expressing FAST in the cytoplasm, the fluorescence from FAST was only observed in the presence of HBR (Figure 4.3). Even after 30 minutes incubation with HBRAA-3M, there was no fluorescence signal from FAST (Figure 4.3), which demonstrated that HBRAA-3M could not enter into cells to label FAST expressed in the cytoplasm of HeLa cells. Theses cell experiments proved that the strategy of introducing a negative charged acetate group on the rhodanine head worked well to design cell impermeant HMBR analogues for FAST and the brightness of FAST:HBRAA-3M was reasonable for imaging FAST-tagged protein in living cells through microscopy.

We next tried to improve brightness of the membrane-impermeant FAST systems by engineering the fluorogen structure. The brightness of FAST:fluorogen complexes depend on the ability of the protein cavity to block the fluorogen in the appropriate emissive conformation. We presented previously that the substituents on the aromatic ring were key determinants for the affinity and brightness of FAST:fluorogen. We thus designed and synthesized a collection of HBRAA-3M analogues bearing different substituents on the phenol ring through condensation reaction of the starting aldehydes and rhodanine acetic acid Switching the methyl group in position 3 to a methoxy group in position 2 of the phenol ring resulted in a compound HBRAA-2OM that bound FAST with higher affinity (Table 4.1).

(Figure 4.2).
However, the resulting complex exhibited reduced fluorescence brightness (Table 4.1). On the other hand, adding an additional methyl group in meta-position of the phenolic hydroxyl group of HBRAA-3M gave a compound HBRAA-2,5DM resembling HBRAA-3M in terms of affinity and brightness (Table 4.1). Compared to FAST:HBRAA-3M, FAST:HBRAA-2,5DM exhibited a 20nm red shift in absorption and a 10nm red shift in emission (Table 4.1). HBRAA-3,5DOM with two methoxy groups in ortho-positions of the phenolic hydroxyl group bound FAST with really low affinity (K D > 20uM), which was not suitable for FAST.

However, FAST:HBRAA-3,5DM exhibited a 50 nm red shift in absorption and a 65 nm red shift in emission compared to FAST:HMBR, absorbing green light and fluorescing red.

Although it is not a good fluorogen for FAST, it displayed significant potential to engineer red-emitting membrane-impermeant FAST system. The brightest FAST:fluorogen complex was eventually obtained with HBRAA-3E, in which the methyl was replaced by an ethyl group. HBRAA-3E bound FAST with affinities around 1 M. The resulting complex FAST:HBRAA-3E was 50 % as bright as FAST:HMBR (Table 4.1), reaching brightness reasonable for cell imaging. Note that FAST:HBRAA-3E exhibited also a 25 nm red shift in absorption and a 20 nm red shift in emission compared to FAST:HMBR (Table 4.1). Interestingly, replacing the ethyl group by an ethoxy group (HBRAA-3OE) further red-shifted the absorption and the emission. FAST:HBRAA-3OE exhibited a 45 nm red shift in absorption and a 35 nm red shift in emission compared to FAST:HMBR.

To sum up, molecular engineering allowed us to increase the affinity and the fluorescence properties of the membrane-impermeant fluorogens. However, there is still some room for improvement both in terms of brightness and affinity. In consequence, new protein tags should be engineered in the future to optimize the system.

Labeling of surface exposed FAST-tagged proteins in live cell with HBRAA-3E

We next evaluated the ability of HBRAA-3E to label cell-surface exposed FAST-tagged proteins in living cells. We expressed transiently in HeLa cells FAST fused at the N-terminus to the murine Ig -chain leader sequence, which directs the protein to the secretory pathway, and at the C-terminus to the platelet derived growth factor receptor (PDGFR) transmembrane domain, which anchors the protein to the plasma membrane, displaying FAST on the extracellular side. Transfected cells were treated with media containing 5 M of HMBR or HBRAA-3E. Confocal microscopy allowed us to show that HMBR labeled both cell-surface exposed FAST and intracellular FAST still present within the secretory pathway, while HBRAA-3E labeled selectively cell-surface proteins (Figure 4.4), in agreement with HBRAA-3E being membrane impermeant. Addition of membrane-permeant HMBR to cells pre-labeled with HBRAA-3E allowed visualizing intracellular FAST within the secretory pathway (Figure 4.4), further demonstrating that HBRAA-3E did not cross the plasma membrane. Even after two hours incubation with media containing 5 M HBRAA-3E, there is no intracellular labeling (Figure 4.4) observed, allowing anticipating the use of HBRAA-3E for long-term studies. Note that we also verified that long exposure to HBRAA-3E (> 24 h) was non-toxic for mammalian cells (Figure 4.5).

Screening of new protein tags for HBRAA-3,5DOM

The red-shifted spectral properties of HBRAA-3,5DOM inspired us to engineer an optimal protein tag for HBRAA-3,5DOM for multi-color cell-surface protein labeling. With the same methodology used in Section III-2, we selected FAST variants that could tightly bind with HBRAA-3,5DOM and form complexes displaying bright red emission from the library of FAST variants (See Section III-2). We screened our library of yeast surface-displayed FAST variants in the presence of 5 M HBRAA-3,5DOM by high throughput FACS (Figure 4.6).

We selected yeast cells incubated with HBRAA-3,5DOM in the condition of Ex 561 nm/Em 614±20 nm by FACS. The fluorescence of Alexa 647 was detected in the condition of Ex 640 nm/Em 671±30 nm. For the first round of FACS we collected about 5% yeast population to make sure that we collected enough positive yeast cells for enriching fluorogen-activating population. We reduced gradually the collecting percentage of yeast population (second round: 4%, third round: 0.4%). From the fourth round, we observed the existing of a fluorogenactivating cell population. We reduce the collecting percentage of cell population to 0.05% for accumulating fluorogen-activating population efficiently. After 5 FACS rounds we obtained a fluorogen-activating cell population. Individual clones were isolated and tested by flow cytometry. Among 24 selected clones, 22 The spectroscopic data of FAST: HBRAA-3E is given for comparison.

IV-3 Conclusion and perspective

In conclusion, we developed a collection of membrane-impermeant fluorogens that bind The promising applications of cell-impermant FAST approach motivate us to extend its spectral properties. We are selecting FAST variants that could bind new cell impermeant fluorogens forming fluorescent complexes with different colors and improved brightness from the library of FAST variants. After 5 rounds of FACS, 5 FAST variants displaying fluorogenactivation in the presence of HBRAA-3,5DOM have been selected. Three protein variants were purified and spectral properties of variant:HPAR complexes were characterized.

Actually, bright red-emitting membrane-impermeant complexes which displayed a ~55nm red shift in absorption ( abs ~560 nm) and a 56 nm red shift in emission ( em ~615 nm) compared to the previous FAST:HBRAA-3E were obtained, which made the application of cellimpermeant FAST systems more versatile.

The other two variants will later be expressed and then spectral properties of FAST variant:

HBRAA-3,5DOM complexes will be characterized. Variants which are able to activate specifically the red fluorescence of HBRAA-3,5DOM preserving unique feature of FAST approach will be identified and compared with the three previous characterized complexes.

Next the performance of the best red-emitting cell-impermeant complex will be tested in living cells.

IV-4 Materials and Methods

General protocol for the synthesis of the membrane-impermeant fluorogens A solution

containing rhodanine-3-acetic acid (191 mg, 1.0 mmol) and the substituted 4-hydroxybenzaldehyde (1.0 mmol) in 40 mL of water was stirred at 90°C for 7 days. After cooling to 4°C and standing overnight, the precipitate was filtered through a glass filter and the crude solid was washed with water, ethanol and dried over P 2 O 5 , to give the desired product.

(Z)-2-(5-(4-hydroxy-3-methylbenzylidene)-4-oxo-2-thioxothiazolidin-3-yl)acetic acid

1H), 4.73 (s, 2H), 4.10 (q, J = 6.9 Hz, 2H), 1.37 (t, J = 6.9Hz, 3H); [START_REF] Gahlmann | Exploring bavterial cell biology with single-molecule tracking and super-resolution imaging[END_REF] Shoreline, WA). The overall emission quantum yields after one-photon excitation  were determined as previously described [START_REF] Plamont | Small fluorescence-activating and absorption-shifting tag for tunable protein imaging in vivo[END_REF] . Affinity constants were determined by spectrofluorometric titration using a Spark 10M plate reader (Tecan) following protocols previously described [START_REF] Plamont | Small fluorescence-activating and absorption-shifting tag for tunable protein imaging in vivo[END_REF] .

Molecular Biology

The plasmid pAG211 for mammalian expression of FAST (fused also to mCherry as additional transfection marker) at the outer plasma membrane was obtained by inserting the sequence coding the protein of interest within the pDisplay plasmid (TermoFisher) using Sal I and Bgl II restriction sites. The plasmid pAG96 for cytoplasmic expression of FAST fused to mCherry and the plasmid pAG 87 for bacterial expression of FAST were previously reported [START_REF] Plamont | Small fluorescence-activating and absorption-shifting tag for tunable protein imaging in vivo[END_REF] . The plasmids pAG286, pAG287 and pAG290 driving bacterial and in vitro expression of clone 1, clone 2 and clone 5, respectively, with an Nterminal His-tag under the control of a T7 promoter were obtained by inserting the gene encoding clone 1, clone 2 or clone 5 between Nhe I and Xho I restriction sites in the pET28a vector. Flow cytometry analyses were performed on an Accuri C6 cytometer (BD Biosciences).

Bacterial expression and protein purification

Chapter V General discussion

Fluorogen-based systems for specific protein labeling play a more and more significant role in live cell imaging because they enable one to achieve high imaging contrast and to adapt properties or modify it with specific groups to achieve new functional properties.

V-1 Development of new fluorogens

The fluorogen HMBR for FAST is based on a push-pull system. An electron-donating phenol ring contacts with the electron-withdrawing rhodanine ring through a conjugated system. It We report a collection of fluorogens enabling to tune the fluorescence color of FAST from green-yellow to orange and red. Beyond allowing multicolor imaging of FAST-tagged proteins in live cells, these fluorogens enable dynamic color switching because of FAST's reversible labeling, opening great prospects for the design of selective imaging methods relying on dynamic systems. In order to further expand the spectral properties of FAST to red, we also designed and developed a library of red fluorogenic dyes, for which we engineered specific protein binders by applying a directed evolution strategy based on the yeast display technology and high-throughput fluorescence activating cell sorting (FACS). We finally developed novel fluorogens able to form fluorescent complexes with FAST, but incapable of crossing the plasma membrane, which makes it possible to selectively detect FAST-tagged cell-surface proteins.

2

 2

Figure 1 . 1

 11 Figure 1.1 Perrin-Jablonski diagram. S 0 : singlet fundamental electronic state, S 1 , S 2 : singlet excited electronic states, T 1 , T 2 : triplet electronic states, IC: internal conversion, ISC: intersystem crossing. Adapted from Valeur.2

Figure 1 . 2

 12 Figure 1.2 Absorption and emission spectra of fluorescein. Absorption and emission spectra are symmetrical. The gap between the maximum wavelength of absorption and emission spectra is the Stokes shift.

Figure 1 . 3

 13 Figure 1.3 Color regions of visible light. violet: 400-455nm, blue: 455-492nm, green:492-577nm, yellow: 577-597nm, orange: 597-620nm, red: 620-700nm.

Figure 1 . 3

 13 Figure 1.3 Common organic fluorescent dyes with various spectral properties. Adapted from Raines et al.[START_REF] Lavis | Bright ideas for chemical biology[END_REF]

Figure 1 . 4 Figure 1 . 5

 1415 Figure 1.4 Genetic fusion of GFP to a POI enables specific fluorescent protein labeling.

Figure 1 . 6

 16 Figure 1.6 Biological applications of AFPs in scientific research. Adapted from Lukyanov et al.7

29 Figure 1 . 30 I- 2 . 2 . 2

 29130222 Figure 1.7 AFP-based techniques for studying protein-protein interaction. a) FRET: ECFP (donor) transfers its energy to EYFP (acceptor) when two fusion proteins interact. b) BiFC: Interaction between two fusion proteins enables the reconstitution of AFP. P: protein, D: donor, A: acceptor, ECFP: enhanced cyan fluorescent protein, EYFP: enhanced yellow fluorescent protein.Adapted from Frommer et al.[START_REF] Frommer | Genetically encoded biosensors based on engineered fluorescent proteins[END_REF]

Figure 1 . 8

 18 Figure 1.8 The formation of the chromophore HBI in the binding barrier of GFP. This multi-step maturation process requires the presence of O 2 and takes tens of minutes or hours.

Figure 1 .

 1 Figure 1.9 a) Chemical stucture of biarsenical substrates FlAsH-EDT 2 and ReAsH-EDT 2 . b) Schematic illustration of the biarsenical-tetracysteine tagging system. FlAsH as an adduct with two 1,2-ethanedithiol (EDT) is weakly fluorescent but becomes fluorescent after interacting with thiol groups of the tetracysteine tag POI: protein of interest.

Figure 1 .

 1 Figure 1.10 General mechanism of site-specific labeling systems. SNAP-tag, CLIP-tag and Halotag. POI: Protein of interest.

  are non-fluorescent by their own but become strongly fluorescent only upon specific binding to a given target. The application of fluorogens minimizes unspecific fluorescence background in cells as free fluorogens are almost invisible even in excess 50 and therefore saves the need for washing steps. The simplified labeling protocol and the high imaging contrast have recently placed fluorogen-based reporters to the central spot of fluorescence imaging. Komatsu et al. presented some fluorescence activation-coupled protein labeling (FAPL) probes for the SNAP-tag protein based on Förster resonance energy transfer (FRET)principle.[START_REF] Komatsu | Real-time measurements of protein dynamics using fluorescence activation-coupled protein labeling method[END_REF] They designed SNAP-tag labeling probes with different fluorophores at the benzyl moiety and a quencher at the guanine moiety via a linker attached at the C-8 position of the guanine. Before labeling, these substrates show almost no fluorescence because of the high FRET efficiency between the fluorophore and the quencher. However, after reaction with SNAP-tag, fluorescence rapidly appears as the fluorophore is separated from the quencher leaving with the guanine moiety (Figure1.11). A fluorescence activation ratio of over 300 is observed after labeling, achieving high contrast imaging and avoiding the need of washout process. Komatsu et al showed that the fluorogen-based SNAP-tag system could be used to study real-time protein activities with high temporal resolution.[START_REF] Komatsu | Real-time measurements of protein dynamics using fluorescence activation-coupled protein labeling method[END_REF] Sun et al. further developed intramolecularly quenched fluorogenic probes for SNAP f , a faster labeling variant of SNAP-tag. SNAP f displayed up to tenfold increase in activity towards benzylguanine substrates compared to SNAP26m, a commercially available version of SNAPtag. By exploring different combinations of fluorophore/quencher pairs, they obtained optimal intramolecularly quenched substrates with fluorescence emission across the visible spectrum.[START_REF] Sun | Development of SNAP-tag fluorogenic probes for wash-free fluorescence imaging[END_REF] Except for these intramolecularly quenched fluorogenic probes, the group of Johnsson developed near-infrared fluorogenic silicon-rhodamine (SiR) probes for SNAP-tag, CLIP-tag and Halo-tag (Figure1.11).[START_REF] Lukinavičius | A near-infrared fluorophore for live-cell superresolution microscopy of cellular proteins[END_REF] These SiR probes are environment-sensitive SiR-carboxyl derivatives. In relative polar environment, the carboxylate group of these probes can attack the electrophilic center of the heterocycle forming the non-fluorescent spirolactone. However the coupling to protein tags favours the formation of the open fluorescent zwitterions. The opening of the cyclized chromophore is the important mechanism for the fluorogenic response because of a large change of the absorption properties between the open and closed states (Figure 1.11)

Figure 1 .

 1 Figure 1.11 Fluorogenic site-specific labeling systems. a) Principle of intramolecularly quenched fluorogenic probes for SNAP-tag. Before labeling, fluorescence is quenched because of the high FRET efficiency between the fluorophore and the quencher. After reaction with SNAP-tag protein, the quencher leaves with the guanine moiety and fluorescence appears rapidly. b) Structures and mechanism of near-infrared fluorogenic SiR probes. SiR probes are sensitive to the environment. In relative polar environment, the carboxylate group of SiR probes attacks the electrophilic center of the heterocycle forming the non-fluorescent spirolactone. However coupling to protein tags favours the formation of the open fluorescent zwitterions. POI: protein of interest.

Figure 1 .

 1 Figure 1.12 Structures of fluorogens for FAPs.

Figure 1 .

 1 Figure 1.13 Development of fluorogens for FAPs. a) The DIR based fluorogens which could bind K7 and generate FAP systems with various emission wavelengths. b) Structures of -CN-DIR and fluorinated TO which improve the photostability of FAP system. c) Structures of fluorogens for blue and far-red FAP systems.

Figure 1 .

 1 Figure 1.14 The principle of PYP-tag labeling system. Coumarin thioester derivatives react with PYP through transthioesterification. POI: Protein of Interest.

Figure 1 .Figure 1 . 16

 1116 Figure 1.15 The intramolecular interaction based fluorogen for PYP-tag. a) Principle of fluorogenic labeling of PYP-tag with FCTP. b) Structure of FCANB, a new fluorogen for PYPtag to accelerate the labeling. The cinnamic acid thioester and the introduction of the quencher reduced the intramolecular interaction between ligand and fluorophore moieties and thus enhanced the labeling kinetic.

Figure 1 .

 1 Figure 1.17 Structures of retinoic acids and synthetic retinoids that shows binding affinity with CRABPII.

Figure 1 .

 1 Figure 1.18 Fluorogenic labeling of CRABPII-tag with merocyanine aldehyde.

Figure 1 .

 1 Figure 1.19 The Spinach labeling system. The non-fluorescent DFHBI reacts with the RNA aptamer Spinach and forms a fluorescent complex. Adapted from Jaffrey et al.[START_REF] Strack | Using Spinach-based sensors for fluorescence imaging (HBRAA-3M) Yellow powder (80 %)[END_REF]

Figure 1 . 79 I- 2 . 8

 17928 Figure 1.20 General procedure for isolating fluorescent "light up" aptamers. Millions of aptamers were expressed in Escherichia coli. and selected through FACS (Fluorescence Activated Cell Sorting) in the presence of potential fluorophores. SELEX (Systematic Evolution of Ligands by Exponential Enrichment) Adapted from Jaffrey et al.[START_REF] Filonov | Rapid selection of an RNA mimic of green fluorescent protein by fluorescence-based selection and directed evolution[END_REF]

Figure 1 .

 1 Figure 1.21 Principle of FAST.

(

  HC), binds to its residue Cys69 and shares structure similarity with HMBR. This is an important reason why PYP was chosen as protein scaffold to develop FAST. The engineering of FAST used a fluorescence activating cell sorting (FACS)-based directed evolution strategy (Figure1.22). At first, libraries of PYP mutants were built by saturation mutagenesis of positions in the chromophore binding sites. Yeast-displayed libraries expressing PYP mutants were incubated with the fluorogen and sorted by high throughput FACS.

Figure 1 .

 1 Figure 1.22 Engineering of FAST. Libraries of PYP mutants are built by saturation mutagenesis of loops (the loop 94-101 is given in orange) within the chromophore binding site. Through surface display technology, libraries of yeast cells expressing protein variants on their surface are obtained. Yeast libraries incubated with the fluorogen are sorted by high throughput FACS. Iterative FACS cycles enable to enrich the population of fluorogen-activating variants.

Figure1. 23

 23 Figure1.23 Chemical structure of HMBR (left). HMBR is based on a push-pull system. Electrondonating phenol interacts with electron-withdrawing rhodanine through a conjugated linker. General structure of HMBR and the arrangement of this thesis. HMBR analogues are fully engineered by modifying its electron-donating part (See Chapter II), conjugated linker (See Chapter III) or electronwithdrawing part (See Chapter IV) for developing novel FAST approaches. ED: electron-donating part, CL: conjugated linker, EW: electron-withdrawing part.

  Dynamic multi-color protein labeling in living cells. Chemical Science (2017) just accepted DOI: 10.1039/C7SC01364G. C. Li, M.-A. Plamont, H. Sladitschek, V. Rodrigues, I. Aujard, P. Neveu, T. Le Saux, L. Jullien & A. GautierII-1 Presentation of article 1FAST is a promising fluorogen-based probe for multiplexed imaging with a lot of attractive advantages. To meet requirements for multiplexed imaging, we need to engineer new systems with various properties. A general strategy for the development of protein labeling systems is to extend their spectral properties and finally cover the whole visible spectrum. The marvelous versatility of molecules-based protein labeling systems enables to easily adjust properties of molecules to the need of the experiment through chemical modulations. So we tried to develop HMBR analogs that could selectively bind with FAST, preserve the main features of FAST but expend its spectral properties. Red-emitting probes are particularly attractive for biological research because the use of longer wavelengths reduces cellular autofluorescence, improves imaging contrast and decreases phototoxicity. One part of my PhD project was focused on the development of red-emitting FAST system. It was shown previously that FAST could accept various substituents on the aromatic ring. Moreover, aromatic substituents are a priori key determinants of the fluorescence efficiency, as they can play a major role in immobilizing the fluorogen within FAST cavity. In this chapter, we explored the role of substituents on the phenol ring of HMBR in varying spectral properties and binding affinities of FAST:fluorogen complexes. New fluorogens with conserved binding ability with FAST and showing different spectral properties are reported.

Figure 2 . 1

 21 Figure 2.1 Structures of new fluorogens for FAST and general strategy for the synthesis of new fluorogens.

Figure S2 .

 S2 Figure S2. Viability assay of HeLa cells incubated for 24 h with solutions of HBR-3,5DM, HBR-3OM and HBR-3,5DOM at 5 M. Cell viability was tested by using calceinAM and EthD1 (LIVE/DEAD® viability/cytotoxicity assay kit). CalceinAM is a cell-permeant profluorophore cleaved by intracellular esterases releasing the green fluorescent polyanionic calcein in live cells. EthD1 (Ethidium homodimer 1) is a non cell-permeant nucleic acid red fluorescent stain that enters only cells with damaged membranes and undergoes a fluorescence enhancement upon binding to nucleic acids, thereby producing a bright red fluorescence in dead cells. Control experiments with HeLa cells non-incubated with dye (A) or incubated for 30 min with 1% hydrogen peroxyde (B) are shown. Cell fluorescence was evaluated by confocal microscopy. The experiment in (C) shows that HBR-3,5DM, HBR-3OM and HBR-3,5DOM are non-toxic for HeLa cells at the concentrations used for imaging.

Figure S3 .

 S3 Figure S3. Dual-color imaging with a single excitation. Confocal micrographs of live HeLa cells co-expressing lyn-EGFP and H2B-FAST labeled with 5 M HBR-3,5DOM upon single excitation at 488 nm (Green emission channel 493-538 nm; Red emission channel 600-797 nm). Scale bars 10 m.

Figure S4 .

 S4 Figure S4. Dynamic color switching in spectrally crowded environment. (A) Time series of live HeLa cells co-expressing lyn-EGFP (membrane), MTS-mCherry (mitochondria) and H2B-FAST (nucleus) upon fluorogen exchange (Green channel Ex/Em 488/493-575 nm; Red channel Ex/Em 543/578-797 nm). Cells were initially stained with 5 M HBR-3,5DOM. Fluorogen exchange was induced by addition of an excess of HMBR at t = 0 s. The final concentrations of HBR-3,5DOM and HMBR were respectively 0.83 M and 4.2 M. See also Movie S2. (B) Temporal evolution of the green and red nuclear fluorescence intensities.

  Chemistry 14, 9253-9261 (2016). C. Li, M.-A. Plamont, I. Aujard, T. Le Saux, L. Jullien & A. Gautier

Figure 3 . 1

 31 Figure 3.1 Structures of the two series of putative fluorogens.

1 III- 1

 11 Before engineering protein scaffolds enabling to bind and activate these new fluorogens, we studied the behavior of these putative fluorogens in mammalian cells by fluorescence microscopy. Appropriate fluorogens are supposed to exhibit no toxicity for cells and barely fluoresce in culture medium and cells. Cytotoxicity test in HeLa cells allowed the identification of the concentration range ensuring no toxicity (See Article 2-Table 3). When imaging HeLa cells incubated with culture media containing 1 M of dye by fluorescence microscopy, we observed two trends in function of the series used. Fluorogens of series Molecular engineering of red emitting fluorogens 68 were invisible in cells as desired (See Article 2-Figure 7), while the fluorogens of series 2 fluorescently labeled specific organelles in cells, as verified by colocalization experiments with commercial organelle trackers (See Article 2-Figure 7 and Figure 8). Although this latter series is not suitable for the development of new FAST systems, it could be applied as specific organelle trackers. Then we tested the binding abilities of the fluorogens in series 1 with FAST. Except for 1c which displayed a little fluorescence activation, the other fluorogens did not show any detectable binding (by measurement of absorption or fluorescence) to FAST. The larger size of the new fluorogens compared to HMBR is very likely the main raison preventing the binding to FAST. Therefore, we considered engineering complementary protein tags able to accept the newly developed fluorogens.

FAST

  was originally engineered by directed evolution. The high throughput method we chose to select desired variants is the fluorescence-activated cell sorting (FACS). FACS is a cell separating instrument based on fluorescence which was developed for sorting specific cell types from cell mixture. Cell sample should be prepared as fluorescently stained single cell suspension. Under certain pressure, cell suspension is pressed into flow chamber where cells are enclosed by sheath fluid (simple buffer under pressure), leading to a single-cell stream. This stream of cells passes through detection lasers and desired cells emitting fluorescence upon the excitation of laser light were detected by the detector. At the break off point, the stream of cells will be charged accordingly and be partitioned into droplets. Droplets are then separated by passing within an electrical field. Sorting up to 50,000 cells per second, FACS allows one to achieve high throughput cell analysis. As the master factor of the sorting, the fluorescence of cells is usually generated by immunofluorescent labeling with fluorescent dye-tagged antibodies or fluorescent stains. We used the yeast surface display technique to build yeast libraries expressing protein variants on the surface, so desired protein variants could bind with fluorogens in solution and activate their fluorescence. The yeast surface display method was established by the group of Wittrup 2 and has been widely applied for efficient protein engineering by FACS.[START_REF] Plamont | Small fluorescence-activating and absorption-shifting tag for tunable protein imaging in vivo[END_REF] Generally, protein variants are fused to one subunit of a-agglutinin (Aga2p) that is tethered to yeast cell wall by linking with Aga1p (the other subunit of a-agglutinin) through two disulfide bonds (Figure3.2). Usually the hemagglutinin (HA) or c-Myc epitope is fused to the N or C terminal of fusion protein as the site for immunolabeling (Figure 3.2).

Figure 3 . 2

 32 Figure 3.2 Schema of yeast surface display. Protein variants are expressed on the surface of yeast cell as Aga2p fusion protein. Aga2p is attached to yeast cell wall by linking with Aga1p through two disulfide bonds. Fused to the N or C terminal of fusion protein, HA or c-Myc epitope acts as the site of immunolabeling for detecting protein expression. Inspired by Chao et al. 4

  collected fluorogen-activating cell populations that fluoresced in the region of 620±29 nm by FACS. The fluorescence of Alexa 647 was detected in the condition of Ex 640 nm/Em 671±30 nm. For the first round of FACS we collected about 5% of the yeast population to make sure that enough positive yeast cells were collected for enriching fluorogen-activating population. We reduced gradually the collecting percentage of yeast population (second round: 1%, third round: 0.03%). From the fourth round, we observed clearly the presence of a fluorogen-activating population (Figure3.3). We reduce the collecting percentage of population to 0.015% for accumulating fluorogen-activating population efficiently. After 5 rounds of FACS we obtained a fluorogen-activating population. Individual clones were isolated and tested by flow cytometry. Among 24 selected clones,23 clones displayed fluorescence in the presence of HPAR. Genes of the 23 clones were extracted and sequenced. Sequencing revealed only 4 clones: clone 1 (appeared 19 times), clone 2, clone 3 (appeared 2 times) and clone 4 (Figure 3.4).

Figure 3 . 3

 33 Figure 3.3 Graphic data of 5 FACS rounds for engineering protein tag in presence of HPAR. Histograms of the distribution of population exhibiting different fluorescence intensity in the condition Ex 488 nm/Em 620±29 nm.

Figure 3 . 4

 34 Figure 3.4 Sequence alignment of the four selected clones with FAST

Figure 3 . 5 Table 3 . 1

 3531 Figure 3.5 Absorption (left) and emission (right) spectra of free HPAR and protein variants:HPAR complexes in PBS pH 7.4 at 25 °C.

Figure 3 . 6

 36 Figure 3.6 Comparison of absorption and emission of new protein variants:HPAR complexes with FAST:HBR and FAST:HBR-3,5DOM. The spectra were recorded in PBS pH7.4 at 25 °C.

Figure 4 . 1

 41 Figure 4.1 Chemical structure of HBR and HBRAA

Figure 4 . 2

 42 Figure 4.2 Chemical Structure of cell impermeant fluorogens for FAST and general strategy for the synthesis.

Figure 4 . 3

 43 Figure 4.3 Confocal micrographs (left) of yeast cells expressing Aga2p-FAST incubated with 20 μM HBR or HBRAA-3M. The Aga2p fusion protein was immunolabeled with an Alexa647-

Figure 4 . 4

 44 Figure 4.4 Selective labeling of cell-surface FAST-tagged proteins with membrane-impermeant fluorogens. (a-b) Confocal micrographs of HeLa cells expressing FAST on the extracellular side. (a) Cells were imaged after 5 minutes of treatment with 5 M HMBR or HBRAA-3E. While HMBR revealed membrane and intracellular proteins, HBRAA-3E revealed only surface-exposed proteins. (b) Subsequent addition of cell permeant HMBR and immediate imaging (< 10 s) revealed proteins within the secretory pathway. (c) Confocal micrographs of HeLa cells expressing FAST fused to the red fluorescent protein mCherry treated with 5 M HMBR for 15 seconds or 5 M HBRAA-3E for 2 hours. Side-by-side images were recorded using the same settings for direct comparison of the fluorescence intensities. (a-c) Scale bars 10 m.

Figure 4 . 5

 45 Figure 4.5 Viability assay of HeLa cells incubated for 24 h with solutions of HBRAA-3E at 5 M. Cell viability was tested by using calceinAM and EthD1 (LIVE/DEAD® viability/cytotoxicity assay kit). CalceinAM is a cell-permeant profluorophore cleaved by intracellular esterases releasing the green fluorescent polyanionic calcein in live cells. EthD1 (Ethidium homodimer 1) is a non cellpermeant nucleic acid red fluorescent stain that enters only cells with damaged membranes and undergoes a fluorescence enhancement upon binding to nucleic acids, thereby producing a bright red fluorescence in dead cells. Control experiments with HeLa cells non-incubated with dye (the first row) or incubated for 30 min with 1% hydrogen peroxyde (the second row) are shown. Cell fluorescence was evaluated by confocal microscopy. The experiment (in the last row) shows that HBRAA-3E are non-toxic for HeLa cells at the concentrations used for imaging.

  clones displayed fluorescence activation in the presence of HBRAA-3,5DOM. The genes of the 22 clones were extracted and sequenced. The sequencing revealed five single clones: clone 1 (appeared 16 times) clone 2, clone 3 (appeared 2 times) clone 4 and clone 5 (appeared 2 times) (Figure 4.7).

Figure 4 . 6

 46 Figure 4.6 Graphic data of 5 FACS rounds for engineering protein tag in presence of HBRAA-3,5DOM. Histograms of the distribution of population exhibiting different fluorescence intensity in the condition Ex 561 nm/Em 614±20 nm.

Figure 4 . 7

 47 Figure 4.7 Sequence alignment of the five selected clones with FAST

Figure 2

 2 Figure 4.8 Absorption (left) and emission (right) spectra of free HBRAA-3,5DOM and protein variants:HBRAA-3,5DOM complexes in PBS pH 7.4 at 25 °C.

Figure 4 . 9

 49 Figure 4.9 Comparison of absorption and emission of new protein variants: HBRAA-3,5DOM complexes with FAST: HBRAA-3E. The spectra were recorded in PBS pH7.4 at 25 °C.

FAST and give fluorescent

  complex with various spectral properties. Their inability to cross the plasma membrane enables to selectively image surface exposed FAST-tagged proteins, without labeling intracellular proteins present in the secretory pathway. It shows potential to obtain spatial information on FAST-tagged proteins without the need of microscopy and image analysis by using fluorescence-based techniques such as spectrofluorimetry and flow cytometry, opening great prospects for the design of high throughput screening assays for the discovery of drugs to treat trafficking related diseases.

  Beads were washed with 20 volumes of PBS containing 20 mM Imidazole, and with 5 volumes of PBS complemented with 40 mM Imidazole. His-tagged proteins were eluted with 5 volumes of PBS complemented with 0.5 M Imidazole, followed by dialysis with PBS.Mammalian Cell Culture HeLa cells were cultured in DMEM supplemented with phenol red, Glutamax I, 10% (vol/vol) fetal calf serum (FCS), and 1% penicillin-streptomycin at 37 °C within a 5% CO 2 atmosphere. For microscopic imaging, cells were seeded in μDish or μSlide IBIDI (Biovalley) coated with poly-L-lysine. Cells were transiently transfected using Genejuice (Merck) according to the manufacturer's protocol. Before imaging, cells were washed with PBS, and treated with DMEM media (without serum and phenol red) containing the fluorogens at the indicated concentration. Cells were imaged directly without washing.Cell viability assayHeLa cells were treated with DMEM media containing the fluorogens at the indicated concentrations for the indicated times. Cell viability was evaluated by fluorescence microscopy using the LIVE/DEAD ® viability/cytotoxicity assay kit (Molecular Probes, Life Technologies) following the manufacter's protocol. Fluorescence Analysis Confocal micrographs were acquired on a Zeiss LSM 710 Laser Scanning Microscope equipped with a Plan Apochromat 63×/ 1.4 NA oil immersion objective. ZEN software was used to collect the data. Images were analyzed with Image J.

  experimental requirement by simple chemical modification of the fluorogens. The development of FAST has expanded the application of fluorogen-based specific labeling systems with a lot of impressive advantages. A unique feature of FAST is its highly tunable labeling. Its fluorescence can be switched on or off rapidly by addition or removal of the fluorogen, which opens new opportunities for multiplexed imaging. My PhD aimed to enlarge the FAST family by developing novel systems with different properties for multiplexed imaging. This thesis presents general strategies used for engineering hybrid systems composed of a fluorogenic chromophore and a protein tag. As this type of system contains two parts, the molecule and the genetically encoded tag, the engineering of new systems obviously has two directions, chemical modification of the molecule and engineering of the tag. For chemical modification, one can change functional groups to tune its spectral

  deexcites non-radiatively in solution but relaxes radiatively to the ground state in FAST protein cavity. Based on this push-pull system, we have developed several series of HMBR analogues by changing substituents on the phenol ring, by elongating the conjugated system or by modifying the rhodanine head to design novel fluorogens suitable for protein labeling. I have explored the role of substituents on the phenol ring of HMBR for varying spectral and affinity properties of FAST:fluorogen complexes. Aromatic substituents are a priori key determinants of the fluorescence properties, as they can play a major role in tuning V-1 Development of new fluorogens 120 absorption/emission properties and immobilizing the fluorogen within FAST cavity. New fluorogens preserving the binding ability with FAST and displaying different spectral properties are reported. Notably, orange-emitting FAST:HBR-3,5DM is 2.5-fold brighter than FAST-HMBR, reaching the brightness level of enhanced green fluorescent protein. Furthermore, a red-emitting FAST:fluorogen complex displaying comparable brightness as the red fluorescent protein mCherry was obtained using HBR-3,5DOM as fluorogen. It expands the application of FAST approach because one can adapt the color of FAST to different spectral conditions. Particularly, the possibility to perform dynamic color switching (red-green) by changing the fluorogen solution (HBR-3,5DOM -HMBR) opens a new dimension for multiplexed imaging. One can use the kinetic signature provided by the color exchange to extract the signal of FAST in the presence of both green and red fluorescent proteins, even though spectral crowding prevents spectral discrimination. As the red-emitting fluorescent reporters interests biologists specially, I next elongated the conjugation system of the fluorogen by adding an additional double bound between the phenol ring and the electron-withdrawing heterocycle to develop far-red-emitting FAST systems. As their relatively large size prevented their binding to FAST, I selected FAST variants able to bind HPAR (one of the far-red fluorogens) and activated its far-red fluorescence (vide infra). New protein tags that can bind tightly with new red-emitting fluorogens and specifically activate their fluorescence are thus required. Finally, I developed cell-impermeant fluorogens by adding a negative charged acetate group in the rhodanine head of HMBR. After additional engineering of the phenol group, HBRAA-3E was chosen as the fluorogen for selective imaging of FAST-tagged cell-surface proteins in living cells because its superior fluorogenic performance and affinity. HBRAA-3E enabled to selectively image surface exposed FAST-tagged proteins, without labeling intracellular proteins present in the secretory pathway. This approach displays the potential to obtain spatial information on FAST-tagged proteins by using fluorescence-based techniques such as spectrofluorimetry and flow cytometry, opening great prospects for the design of high Résumé L'étude de la dynamique des protéines est essentielle pour comprendre les processus biologiques. Notre laboratoire a développé une nouvelle classe de protéines fluorescentes semi-synthétiques, appelée Fluorescence-Activating and absorption-Shifting Tag (FAST). Cette thèse de doctorat présente le développement de nouveaux systèmes FAST avec diverses propriétés pour l'imagerie multiplexée. Nous avons développé une série de fluorogènes permettant de modifier la couleur de FAST de vert-jaune à orange et rouge. Au delà de l'application de l'imagerie multi-couleurs, ces fluorogènes permettant un échange dynamique des couleurs grâce à la liaison réversible de FAST, ouvrant de nouvelles perspectives pour le développement de méthodes d'imagerie sélective reposant sur la dynamique de systèmes réactifs. Pour étendre davantage les propriétés spectrales de FAST vers le rouge lointain, nous avons développé une nouvelle série de fluorogènes rouges, pour lesquels nous avons sélectionné par une stratégie d'évolution dirigée basée sur le yeast display et la cytométrie en flux de nouveaux tags protéiques capables d'interagir avec ces fluorogènes et d'activer leur fluorescence.Nous avons enfin développé de nouveaux fluorogènes capables de former des complexes fluorescents avec FAST, mais incapables de traverser la membrane plasmique, ce qui permet de détecter , a small fluorogen-based reporter enabling to fluorescently label fusion proteins in living cells. My PhD thesis presents the developments of new FAST systems with various properties for multiplexed imaging.

  ). At first, a small region of interest (ROI) is irradiated by intense light to bleach AFPs in this region. Then the rate of fluorescence recovery in this region is measured, which reflects the rate of migration of unbleached AFPs from nearby regions, so that one can estimate mobility of AFP-

	labeled biomolecules.
	There is another related strategy, fluorescence loss after photobleaching (FLIP). Instead of
	bleaching ROI, one can bleach a region nearby ROI and measure the rate of fluorescence loss
	of ROI, which avoid influence of possible photodamage in the observed region.

Actually, Photobleaching technique suffers from some inevitable disadvantage: phototoxicity caused by intense light irradiation; photobleaching occuring during the measurement of fluorescence recovery (loss); influence of AFP influx to ROI during bleaching. However, it has been well established because it is simple and straightforward to operate and generally it can provide valuable information on the mobility of biomolecules.

  Zanotti K.J., Silva G.L., Creeger Y., Robertson K.L., Waggoner A.S., Berget P.B.,

	Armitage B.A., Blue fluorescent dye-protein complexes based on fluorogenic cyanine dyes
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	activating proteins, ACS Chem. Biol., 2015, 10, 1239-1246.
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	67 Kleywegt G.J., Bergfors T., Senn H., Le Motte P., Gsell B., Shudo K. Jones T.A., Crystal
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	acid and a synthetic retinoid, Structure, 1994, 2, 1241-1258.
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18, 1473-1483 (2017). C. Li, A. G. Tebo & A. Gautier I-4 References 1 Weissleder R., Nahrendorf M., Advancing biomedical imaging, Proc. Natl. Acad. Sci. U. S. A., 2015, 112, 14424-14428. 2 Valeur B., Molecular Fluorescence: Principles and Applications, Wiley-VCH, 2001. 3 Valeur B., Berberan-Santos M.N., A brief history of fluorescence and phosphorescence before the emergence of quantum theory, J. Chem. Educ., 2011, 88, 731-738. 4 Stokes G.G., Philos. Trans., 1852, 142, 463-562. 5 Reichman J., Handbook of optical filters for fluorescence microscopy, Chroma Technology Corp., 1998. 6 Lakowicz J.R., Principles of Fluorescence Spectroscopy, 3 rd edition, Springe, 2006. 7 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. 8 Demchenko A.P., Introduction to fluorescence sensing, Springer Science + Business Media B.V., 2009. 58 61 Hori Y., Nakaki K., Sato M., Mizukami S., Kikuchi K., Development of protein-labeling probes with a redesigned fluorogenic switch based on intramolecular association for no-wash live-cell imaging, Angew. Chem. Int. Ed. Engl., 2012, 51, 5611-5614. 62 Hori Y., Norinobu T., Sato M., Arita K., Mizukami S., Kikuchi K., Development of fluorogenic probes for quick no-wash live-cell imaging of intracellular proteins, J. Am. Chem. Soc., 2013, 135, 12360-12365. 63 Hori Y., Hirayama S., Sato M. Kikuchi K., Redesign of a fluorogenic labeling system to improve surface charge, brightness, and binding kinetics for imaging the functional localization of bromodomains, Angew. Chem. Int. Ed. Engl., 2015, 54, 14368-14371. 64 Kamikawa Y., Hori Y., Yamashit K., Jin L., Hirayama S., Standleyb D.M., Kikuchi K., Design of a protein tag and fluorogenic probe with modular structure for live-cell imaging of intracellular proteins, Chem. Sci., 2016, 7, 308-314. 65 Hirayama S., Hori Y., Benedek Z., Suzuki T., Kikuchi K., Fluorogenic probes reveal a role of GLUT4 N-glycosylation in intracellular trafficking, Nat. Chem. Biol., 2016, 12, 853-859. 66 Meyer T.E., Isolation and characterization of soluble cytochromes, ferredoxins and other chromophoric proteins from the halophilic phototrophic bacterium Ectothiorhodospira halophila, Biopchim. Biophys. Acta, 1985, 806, 175-183. Rev. Nutr., 1996, 16, 205-234. absorption-shifting tag for tunable protein imaging in vivo, Proc. Natl. Acad. Sci. U. S. A., 2016, 113, 497-502.

Table 3 .2 Physicochemical properties protein variants:HPAR complexes in PBS pH

 3

		Brightness	Stokes shift
	%		nm

7.4. The abbreviations are as follows:  abs , the wavelength of maximal absorption;  em , the wavelength of maximal emission; , the molar absorption coefficient at  abs ; , the fluorescence quantum yield. The spectroscopic data of FAST:HBR and FAST: HBR-3,5DOM are given for comparison.

  Cells were then washed with 10 mL DPBS-BSA, and incubated in 200 μL of 1/100 secondary antibody Alexa Fluor® 647-goat anti-rabbit IgG (Life Technologies) solution in DPBS-BSA for 20 min on ice. After washing with DPBS, cells were incubated in 10 mL DPBS supplemented with 10 μM HPAR, and sorted on a MoFlo™ Astrios Cell Sorter (Beckman Coulter) equipped with a 488 nm and a 640 nm laser.

	9 cells) was grown overnight (30°C, 280 rpm) in 1 L of SD (20 g/L
	dextrose, 6.7 g/L yeast nitrogen base, 1.92 g/L yeast synthetic dropout without tryptophane,
	7.44 g/L NaH 2 PO 4 and 10.2 g/L Na 2 HPO 4 -7H 2 O, 1% penicillin-streptomycin 10,000 U/mL).
	Yeast culture was diluted to OD 600 1 in 1L of SD and grown (30°C, 280 rpm) until OD 600 2-5.
	5 × 10 9 cells yeast cells were then collected and grown for 36 h (23°C, 280 rpm) in 1L SG (20
	g/L galactose, 2 g/L dextrose, 6.7 g/L yeast nitrogen base, 1.92 g/L yeast synthetic dropout
	without tryptophane, 7.44 g/L NaH 2 PO 4 and 10.2 g/L Na 2 HPO 4 -7H 2 O, 1% penicillin-
	streptomycin 10,000 U/mL). 5× 10 8 induced cells were then pelleted by centrigugation (25°C,

3 min, 2,500 g), washed with 10 mL DPBS-BSA (137 mM NaCl, 2.7 mM KCl, 4.3 mM Na 2 HPO 4 , 1.4 mM KH 2 PO 4 , 1 g/L bovine serum albumin, pH 7.4), and incubated for 30 min at room temperature in 200 μL of 1/250 primary antibody chicken antic-Myc IgY (Life Technologies) solution in DPBS-BSA.

  Table 4.1). The loss of affinity and brightness resulted very likely from steric clashes between the added acetate group and FAST.

	Complex	 abs nm	 em nm	 mM -1 cm -1	 %	Brightness	K D M
	FAST:HMBR	481	540	45	23	10,300	0.13
	FAST:HBRAA-3M	502	557	56	6	3,400	6.4
	FAST:HBRAA-2OM	497	537	55	5	2,800	0.48
	FAST:HBRAA-2,5DM	519	566	63	5	3,200	5.4
	FAST:HBRAA-3E	505	559	61	8	4,900	1.3
	FAST:HBRAA-3OE	525	575	62	6	3,700	1.7
	FAST:HBRAA-3,5DOM	532	606				>20

Table 4 .1 Physico-chemical properties of FAST:fluorogen complexes in PBS pH 7.4.

 4 Abbreviations are as follows:  abs , wavelength of maximal absorption;  em , wavelength of maximal emission; , molar absorption coefficient at  abs ; , fluorescence quantum yield; K D affinity constant.

Table 4 .3 Physicochemical properties protein variants:HBRAA-3,5DOM complexes in PBS pH 7.4.

 4 The abbreviations are as follows:  abs , the wavelength of maximal absorption;  em , the wavelength of maximal emission; , the molar absorption coefficient at  abs ; , the fluorescence quantum yield.

Chemical synthesis -general information

  Commercially available reagents were used as starting materials without further purification. NMR spectra were recorded on a AC Bruker spectrometer at 300 MHz for 1 H and 75 MHz for[START_REF] Gahlmann | Exploring bavterial cell biology with single-molecule tracking and super-resolution imaging[END_REF] C; chemical shifts are reported in ppm with protonated solvent as internal reference 1 H, CHCl 3 in CDCl 3 7.26 ppm, CHD 2 SOCD 3 in CD 3 SOCD 3 2.50 ppm;[START_REF] Gahlmann | Exploring bavterial cell biology with single-molecule tracking and super-resolution imaging[END_REF] C,[START_REF] Gahlmann | Exploring bavterial cell biology with single-molecule tracking and super-resolution imaging[END_REF] CDCl 3 in CDCl 3 77.0 ppm,[START_REF] Gahlmann | Exploring bavterial cell biology with single-molecule tracking and super-resolution imaging[END_REF] CD 3 SOCD 3 in CD 3 SOCD 3 39.52 ppm; coupling constants J are given in Hz. Mass spectra (chemical ionization or high resolution) were performed by the Service de Spectrométrie de Masse de Chimie ParisTech and the Institut de Chimie Organique et Analytique de l'Université d'ORléans. Column chromatography was performed on silica gel 60 (0.040-0.063 nm) Merck. Analytical thinlayer chromatography (TLC) was conducted on Merck silica gel 60 F254 precoated plates.

	Synthesis of 4-hydroxy-3-ethylbenzaldehyde To a solution of 2-ethylphenol (6.1 g, 50
	mmol) in 10% aqueous sodium hydroxide (80 mL, 200 mmol) was added trichloromethane
	(15.0 g, 125 mmol) dropwise at 60°C over 1 h, and then the reaction mixture was stirred for 2
	h at 60 ℃. After cooling, the mixture was neutralized by an aqueous solution of hydrochloric
	acid and extracted with dichloromethane. The combined organic layers were washed with
	brine, dried over magnesium sulfate, and concentrated under reduced pressure. The residue
	was purified by flash chromatography on silica gel with cyclohexane/ethylacetate (7.5/2.5, v/v)
	to yield the desired 4-hydroxy-3-ethylbenzaldehyde (1.1 g, 15% yield) as a gray pink solid. 1 H
	NMR (300 MHz, CDCl 3 ,  in ppm): 9.84 (s, 1H), 7.72 (s, 1H), 7.65 (d, J = 8.1 Hz, 1H), 6.92
	(d, J = 8.1 Hz, 1H), 6.51 (s, 1H), 2.70 (q, J = 7.5 Hz, 2H), 1.27 (t, J = 7.5 Hz, 3H); 13 C NMR
	(75 MHz, CDCl 3 ,  in ppm): 192.3, 160.5, 131.5, 131.4, 130.6, 129.4, 115.6, 22.8, 13.5; MS
	(ESI): m/z 149.2[M-H] -, calcd mass for [C 9 H 9 O 2 ] -: 149.1; HRMS (ESI): m/z 149.0608 [M-H] -,
	calcd mass for [C 9 H 9 O 2 ]

-: 149.0603.

  C NMR (75 MHz, CD 3 SOCD 3 ,  in ppm) : 193.0, 167.3, 166.4, 150.7, 147.3, 135.0, 125.5, 124.3, 117.2, 116.5, 115.7, 64.0, 45.0, 14.6; MS (ESI): m/z 338.3 [M-H] -, calcd mass for [C 14 H 12 NO 5 S 2 ] -: 338.0;

	HRMS (ESI): m/z 338.0162 [M-H] -, calcd mass for [C 14 H 12 NO 5 S 2 ] -: 338.0157.
	(Z)-2-(5-(4-hydroxy-3,5-dimethoxybenzylidene)-4-oxo-2-thioxothiazolidin-3-yl)acetic
	acid (HBRAA-3,5DOM) Orange powder (59 %). 1 H NMR (300 MHz, CD 3 SOCD 3 ,  in ppm):
	9.63 (s, 1H), 7.82 (s, 1H), 6.97 (s, 2H), 4.73 (s, 2H), 3.85 (s, 6H); 13 C NMR (75 MHz,
	CD 3 SOCD 3 ,  in ppm) : 193.4, 167.8, 166.8, 148.8(2C), 140.2, 135.7, 123.6, 118.1, 109.3(2C),
	56.6(2C), 45.4; MS (ESI): m/z 354.5 [M-H] -, calcd mass for [C 14 H 12 NO 6 S 2 ] -: 354.0.
	Physical Chemistry experiments pH measurements were performed on a standard pH meter
	PHM210 Radiometer Analytical (calibrated with aqueous buffers at pH 4 and 7 or 10) with a
	Crison 5208 Electrode (Barcelona, Spain). UV/Vis absorption spectra were recorded in 1 cm

× 1 cm quartz cuvettes (Hellma) on a diode array UV/Vis spectrophotometer (Evolution array, Thermo Scientific). Corrected fluorescence spectra upon one-photon excitation were recorded with a Photon Technology International QuantaMaster QM-1 spectrofluorimeter (PTI, Monmouth Junction, NJ) equipped with a Peltier cell holder (TLC50, Quantum Northwest,

  Expression vectors were transformed in Rosetta(DE3)pLysS E. coli (New England Biolabs). Cells were grown at 37°C in Lysogeny Broth (LB) medium complemented with 50 μg/ml kanamycin and 34 μg/ml chloramphenicol °C to allow DNA digestion by DNase. Cellular fragments were removed by centrifugation (9,300 × g for 1h at 4°C). The supernatant was incubated overnight at 4°C under gentle agitation with Ni-NTA agarose beads in phosphate buffered saline (PBS) (sodium phosphate 50 mM, NaCl 150 mM, pH 7.4) complemented with 10 mM Imidazole.

	to OD 600nm 0.6. Expression was induced for 4 h by adding isopropyl β-D-1-
	thiogalactopyranoside (IPTG) to a final concentration of 1 mM. Cells were harvested by
	centrifugation (4,300 × g for 15 min at 4°C) and frozen. The cell pellet was resuspended in
	lysis buffer (phosphate buffer 50 mM, NaCl 150 mM, MgCl 2 2.5 mM, protease inhibitor,
	DNase, pH 7.4) and sonicated (5 min at 20 % of amplitude). The lysate was incubated for 2 h
	at 4

(Z)-5-(4-hydroxy-3,5-dimethylbenzylidene)-2-thioxothiazolidin-4-one (HBR-3,5DM)1 H

Chao G., Lau W.L., Hackel B.J., Sazinsky S.L., Lippow S.M., Wittrup K.D., Isolating and engineering human antibodies using yeast surface display, Nat. Protoc., 2006, 1, 755-768.

Gietz R.D., Schiestl R.H., Large-scale high-efficiency yeast transformation using the LiAc/SS carrier DNA/PEG method, Nat Protoc, 2007, 2,38-41.

Article 1-Figure 3). We observed that labeling of FAST could be switched on and off by addition and withdrawing of fluorogen and that this process was repeatable with various fluorogens which demonstrating the high tunability of the FAST technology.

The discovery of red-emitting FAST: HBR-3,5DOM makes the application of FAST system more versatile because now researchers have additional color options for different experimental conditions. In the presence of red fluorescent probes we can achieve multiplexed imaging by applying HMBR to label FAST-tagged proteins. On the contrary, in the presence of green fluorescent probes we can choose HBR-3,5DOM to label FAST-tagged proteins (See Article 1-Figure 2).

Another outstanding feature of FAST: HBR-3,5DOM complex is its high Stokes shift (82nm).

We succeeded to distinguish FAST fusion protein with HBR-3,5DOM from EGFP-tagged protein by using a single excitation source (488nm) and two separate emission regions (See Article 1-Figure S3), which is significant for multiplexed imaging of living cells because one single excitation source simplifies imaging protocols and reduces phototoxicity.

Furthermore the absorption and emission difference between FAST:HMBR and FAST:HBR-3,5DOM is large enough to distinguish them as two different colors through microscopy (See Article 1-Figure 3). Moreover, we examined and proved that it was possible to switch the color (red-green) of FAST dynamically by changing the fluorogen solution (HBR-3,5DOM -HMBR) (See Article 1-Figure 4a). Using this unique feature, we developped an approach to extract the signal of FAST in the presence of both green and red fluorescent proteins by using only the green and the red channels, which was not evident for common fluorescent reporters because spectral discrimination is impossible in these conditions. The S1. Physico-chemical properties of HMBR, HBR-3,5DM, HBR-2,5DM, HBR-3OM, HBR-3,5DOM in aqueous solutions. Abbreviations are as follows : pK A , acidity constant ;  abs,neutral , wavelength of maximal absorption of the protonated state;  abs,anionic , wavelength of maximal absorption of the anionic state.

Fluorogen

Chapter IV Development of cell impermeant fluorogens for the selective imaging of cell surface proteins

This part has been the object of a patent:

Membrane-impermeant fluorogenic chromophores.

Application number : EP17305591.4

A. Gautier, L. Jullien, C. Li, F. Perez

IV-1 Introduction

Cell surface proteins are important for biological research because they play indispensable roles in various biological activities. Specific labeling of cell surface proteins for studying their surface expression, transportation and function is highly required. Fluorogen-based specific labeling systems have significant potential for specifically labeling cell surface protein because one can easily adapt their methods to experimental condition by simply engineering cell impermeant fluorogens. Nowadays the discovery of small molecules able to restore trafficking defects induced by genetic mutations has become highly important in drug design 1 . Specific surface labeling methods have been applied to test the ability of a small molecule to restore normal traffic. However, spatial information of the protein of interest is usually obtained through microscopy and image analysis, limiting thus screening throughput.

Detecting methods without the need for imaging, such as spectrofluorimetry or flow cytometry that allow high throughput analysis, are thus indispensable 2 .

Fluorogens used in fluorogen-based specific labeling system generally cross the cell membrane and enter into the cell through a spontaneous process named diffusion. The rate of simple diffusion of a molecule across the membrane depends on its size and its hydrophobicity. Ordinarily, small hydrophobic molecules diffuse across the cell membrane easily; polar but small and uncharged molecules can also cross the membrane through diffusion; large polar molecules or ions are not cell-permeant. In sum, potential cell impermeant fluorogens should be polar, charged or large in size.

throughput screening assays for the discovery of drugs to treat trafficking related diseases.

V-2 Selection of new protein tags

We used directed evolution as the strategy for selecting new protein tags. Yeast surface display and FACS were applied to achieve high throughput selection. The possible protein scaffolds should contain reasonable cavities for binding fluorogens and activating their fluorescence. The protein library applied in this thesis for selecting new protein tag was a library of FAST variants constructed by random mutagenesis through error-prone PCR. This protein scaffold obviously shows binding affinity with HMBR analogues and was perfect to engineer variants with improved properties. From this library, I have selected novel protein tags activating the fluorescence of the far-red emitting HPAR and the membrane-impermeant red-emitting HBRAA-3,5DOM.

The described library of FAST variants library brings a lot of possibilities for developping new FAST systems. In addition to the two applications mentioned above, protein engineering for improving the spectral and binding properties of FAST:HMBR, FAST:HBR-3,5DOM and FAST:HBRAA-3E is already on-going in the lab. Multiplexed imaging motivates in particular the engineering of FAST variants displaying orthogonal fluorogen binding specificities to achieve orthogonal labeling.

In conclusion, with (i) its unique advantages for live cell imaging, (ii) the possibility of easily engineering new fluorogens through molecular engineering, and (iii) the robustness of the protein evolution protocol, FAST has become a really promising fluorogen-based labeling system, opening new opportunities for various applications in biological imaging, including multiplexed imaging and cellular biosensing.