Abbreviations

bacterial cell suggests that the cytoplasm is a very crowded environment [START_REF] Kumar | Mobility of Cytoplasmic, Membrane, and DNA-Binding Proteins in Escherichia coli[END_REF][START_REF] Ando | Crowding and hydrodynamic interactions likely dominate in vivo macromolecular motion[END_REF][START_REF] Nenninger | Size Dependence of Protein Diffusion in the Cytoplasm of Escherichia coli[END_REF] . Their cytoplasm has often been described as a complex mixture (like a "soup") containing the chromosome and all the macromolecular machineries necessary for DNA replication, gene expression and other metabolic processes. Bacteria reproduce through a binary fission process: bacterial cell mass increases until a new cell wall grows through the center thus dividing the mother cell and forming two daughter cells, each bacterium is the same as their parent cell.

A B Bacteria are the earliest and longest life forms on earth. They can inhabit every corner of the earth: water, soil, trees, human, animals, even nuclear waste [START_REF] Fredrickson | Geomicrobiology of High-Level Nuclear Waste-Contaminated Vadose Sediments at the Hanford Site, Washington State[END_REF] . A gram of soil contains about 10 6 -10 9 bacteria, and a piece of leaf may contains about 10 4 -10 8 bacteria 4 (Figure 1.1B). Bacteria are very common in our daily life. So it is important to do research on this unseen majority. However, until now people don't really understand how this kind of cell is organized to coordinate the different cellular processes. Studying and improved understanding of bacteria can be used for medical research [START_REF] Lusby | Bactericidal Activity of Different Honeys against Pathogenic Bacteria[END_REF][START_REF] Finegold | Recently Described Clinically Important Anaerobic Bacteria: Medical Aspects[END_REF] , water resources [START_REF] Unc | Transport of bacteria from manure and protection of water resources[END_REF][START_REF] Mckeon | Antibiotic resistant gram-negative bacteria in rural groundwater supplies[END_REF] , and food industry [START_REF] Leroy | Lactic acid bacteria as functional starter cultures for the food fermentation industry[END_REF][START_REF] Gram | Food spoilage-interactions between food spoilage bacteria[END_REF] .

Many bacteria are helpful for humans, for example, bacteria can help to digest food [START_REF] Leroy | Lactic acid bacteria as functional starter cultures for the food fermentation industry[END_REF] .

However people are normally scared by bacteria because some pathogens cause disease in humans race, for example, the pathogenic bacteria (e.g. E. coli O157:H7) hidden in food [START_REF] Leyer | Acid adaptation of Escherichia coli O157:H7 increases survival in acidic foods[END_REF] and water [START_REF] Radke | A high density microelectrode array biosensor for detection of E. coli O157:H7[END_REF] or even medical equipment may cause illness in the human body and can become life-threatening [START_REF] Su | Escherichia coli O157: H7 Infection in Humans[END_REF] . Thus, rapid detection of bacterial growth is important for medical diagnostics to identify contamination in a medical setting [START_REF] Barker | Multiple pH measurement during storage may detect bacterially contaminated platelet concentrates[END_REF][START_REF] Palavecino | Detecting bacterial contamination in platelet products[END_REF][START_REF] Washington | Medical Microbiology[END_REF] and in the food-processing industry in order to prevent contamination of food, air, and water [START_REF] Farris | Principles of Bacterial Detection: Biosensors, Recognition Receptors and Microsystems[END_REF][START_REF] Ko | Self-indicating nanobiosensor for detection of 2,4dinitrophenol[END_REF][START_REF] Wang | New Trends in Impedimetric Biosensors for the Detection of Foodborne Pathogenic Bacteria[END_REF] .

The Escherichia coli (E.coli) bacterium is a gram-negative bacterium that is used as a model organism for research (Figure 1.2). E.coli bacteria are surrounded by phospholipid bilayers cell membrane. As prokaryotes, unlike eukaryotic cells, E.coli bacteria don't have membrane-bound organelles in their cytoplasm. The cytoplasm is a complex mixture containing proteins, ribosomes, plasmid, nucleoid, which is the circular genome of the cell bound by regulatory proteins. E.coli bacteria also have flagella which are used for motility.

E.coli is the most widely used and studied model organism in biotechnology and microbiology. The bacterium can be grown and cultured easily and inexpensively in a laboratory setting, and has been intensively investigated for over many years. E.coli cells are typically rod-shaped, and are about 3 µm long and 0.5 µm in diameter 21,[START_REF] Valeur | Molecular Fluorescence[END_REF] . For this research project, I used the E.coli strains K12 BW25113. This strain is adapted to laboratory setting and is not pathogenic. In this project I aim at developing and characterizing new fluorescent materials in order to develop internal and external imaging of E.coli. I also envision detecting growth of bacteria. Thus different fluorescent nanomaterials were synthetized: such as fluorescent nanoparticles (FNPs Chapters 2, 3, 5) and fluorescent polymer chains (FPCs Chapter 4). I will show how these materials target internal and external parts of E.coli to develop bioimaging and biosensing applications of E.coli bacteria. Before going further, I will now introduce what fluorescence is and why I have chosen to detect such signal.

What is fluorescence?

Fluorescence is a radiative deactivation after absorption of a photon at appropriate energy. The emission of another photon with longer wavelength after a very brief time (few ns) is fluorescence. The processes that occur after the absorption of light by a molecule or a material are usually illustrated by the Perrin-Jablonski diagram (Figure 1.3). The Perrin-Jablonski diagram shows the various electronic states of a molecule, such as the ground state (S 0 ), the excited states (S 1 , S 2 …) and the triplet states (T 1 , T 2 …) [START_REF] Valeur | Molecular Fluorescence[END_REF] . A molecule in the ground state (S 0 ) can absorb light and be excited to higher electronic states like singlet electronic states (S 1 , S 2 …). Molecules in the condensed phases rapidly relax to the lowest vibrational level of S 1 , this process is called internal conversion (IC). Molecules in the S 1 state can also undergo a spin conversion into the first triplet state T 1 , such conversion is called intersystem crossing (ISC). Emission from T 1 to the ground state is called phosphorescence. Emission of photons accompanying the S 1 to S 0 relaxation is called fluorescence. The average time the molecule stays in its excited state S 1 before emitting a photon is called fluorescence lifetime.

Examination of the Perrin-Jablonski diagram (Figure 1.3) indicates that the energy of the emission is less than that of absorption. So, emitted fluorescent light has a longer wavelength and lower energy compared with the absorbed light. This phenomenon, known as Stokes shift (Figure 1.4), is due to the energy loss in the fluorescence process. One common reason of the Stokes shift is the rapid decay to the lowest vibrational level of S 1 .

Furthermore, fluorophores usually decay to higher vibrational levels of S 0 (Figure 1.3), resulting in further loss of excitation energy by thermalization of the excess vibrational energy. Besides these effects, fluorophores can display Stokes shifts due to other reasons, such as solvent effects, excited-state reactions, complex formation, and energy transfer [START_REF] Valeur | Molecular Fluorescence[END_REF][START_REF] Lakowicz | Principles of Fluorescence Spectroscopy[END_REF] . In many cases the absorption spectrum is a mirror image of the emission spectrum. This is the mirror image rule which states that electronic transitions are vertical [START_REF] Lakowicz | Principles of Fluorescence Spectroscopy[END_REF] . The fluorescence quantum yield (Φ F ) defines the efficiency of the fluorescence process. It is known as the ratio of the number of emitted photons to the number of absorbed photons.

The quantum yield is generally independent from the excitation wavelength [START_REF]Handbook of Fluorescence Spectra of Aromatic Molecules[END_REF] . This is because the excited molecules normally decay to the lowest vibrational level of S 1 before fluorescence emission [START_REF] Lakowicz | Principles of Fluorescence Spectroscopy[END_REF] .

Fluorescence intensity can be decreased by a wide variety of processes which is called quenching processes (Figure 1 Another important process is the Förster resonance energy transfer (FRET) (Figure 1.5C). This process occurs when the emission spectrum of a fluorophore (donor), overlaps with the absorption spectrum of another molecule (acceptor). The acceptor is designed to be close enough to the donor, and the acceptor does not need to be fluorescent. Fluorescence in life science has been widely used. People use chemical reactions to label biological macromolecules with fluorescent dyes in order to detect these biomolecules.

An ideal fluorescent probe and label for bioimaging and biosensing application should carry out the following requirements:

1) High brightness.

Brightness is the total efficiency of absorption and fluorescence emission of a fluorophore. Brightness of a dye depends on two properties: molar extinction coefficient at the excitation wavelength ε(λ) and fluorescence quantum efficiency, Φ F [START_REF] Braslavsky | Glossary of terms used in photochemistry, 3rd edition (IUPAC Recommendations[END_REF] .

A fluorophore with high molar extinction coefficient describes a high ability of the fluorophore to absorb light. Light absorbing ability of the fluorophore is one parameter to contribute to the brightness. Another parameter is the fluorescence quantum yield, Φ F , high quantum yield means in higher ratio of the number of emitted photons to the number of absorbed photons which resulted in high fluorescence efficiency.

The brightness (B) of a fluorophore is proportional to the molar extinction coefficient at the excitation wavelength ε(λ) and quantum yield (Φ F ), as indicated in the following relationship [START_REF] Braslavsky | Glossary of terms used in photochemistry, 3rd edition (IUPAC Recommendations[END_REF] :

𝐵 = 𝜀(𝜆) × 𝛷 𝐹
For assemblies of fluorophores (e.g. nanoparticles), the brightness is defined as [START_REF] Grazon | Élaboration de nanoparticules fluorescentes à base de BODIPY par polymérisation RAFT en miniémulsion : synthèse, caractérisation et fonctionnalisation de surface[END_REF] :

𝐵 = 𝑁 × 𝜀(𝜆) × 𝛷 𝐹
Where, N: Number of fluorophores contained inside the nanoparticle ε(λ): molar extinction coefficient of the fluorophore at the excitation wavelength Φ F : fluorescence quantum yield of the nanoparticle From the equation above, one can notice that optimal excitation wavelength is also very important to high brightness. In order to achieve the highest brightness, it is important

to excite fluorophore at wavelengths (λ) close to the maximum of the absorption.

2) High photostability.

During practical bioimaging experiments, after some number of excitation-emission cycles by intense light in fluorescence microscopy, almost all fluorophores can be photobleached. Photobleaching occurs because of some photochemical reactions that often involves molecular oxygen [START_REF] Demchenko | Introduction to Fluorescence Sensing[END_REF] .

The photostability of a fluorophore can be affected by its local environment. Low photostability often limits the quality images to a short time scale which may be a serious drawback for kinetics or mapping in bioimaging (e.g. cells, tissues, bacteria…).

3) Optimal solubility, penetration, specificity, nontoxicity

High cell permeability and a good intracellular solubility of the label are important for bioimaging and biosensing. High affinity and high specificity for the target is also needed as well as nontoxic effect especially for in vivo research.

From all the parameters described above, one can notice that the ideal fluorescent labeling material hardly exists. Scientists have to consider about the experimental setting and different parameters to make a compromise for each application [START_REF] Valeur | Molecular Fluorescence[END_REF] . In the next paragraph §1.3, different selected typical fluorescent materials are briefly described, followed by the discussion about biosensing and bioimaging applications of these fluorescent materials (organic dyes, fluorescent proteins, quantum dots, silica nanoparticles, organic nanoparticles…).

Fluorescent probes and labels for bioimaging and biosensing

Sensing and labelling are two important methods that are widely used in life science.

Different techniques and strategies based on different physical principles have been developed, such as: mass difference measurement (microcantilever) [START_REF] Carrascosa | Nanomechanical biosensors: a new sensing tool[END_REF] , electron and matter interaction (electrochemistry) [START_REF] Palecek | New approaches in the development of DNA sensors: hybridization and electrochemical detection of DNA and RNA at two different surfaces[END_REF][START_REF] Warsinke | Electrochemical immunoassays[END_REF] , photon and matter interaction (optical sensors) [START_REF] Baird | Current and emerging commercial optical biosensors[END_REF][START_REF] Homola | Present and future of surface plasmon resonance biosensors[END_REF] , advanced microscopy (TEM [START_REF] Reimer | Transmission Electron Microscopy: Physics of Image Formation and Microanalysis[END_REF] , SEM [START_REF]Principles and techniques of scanning electron microscopy[END_REF] ). These techniques however generally need sophisticated and expensive instrumentation which limits their application [START_REF] Demchenko | Introduction to Fluorescence Sensing[END_REF] .

Fluorescent sensing and labelling is one of the most common methodologies used for bioanalytical purposes. Fluorescence techniques are considered to be very sensitive [START_REF] Demchenko | Introduction to Fluorescence Sensing[END_REF] , since their sensitivity can reach in some cases the limit of single molecule. Fluorescence techniques are also considered to be very fast, the response time depends on the fluorescence lifetime which can be about 10 -8 -10 -10 s [START_REF] Demchenko | Introduction to Fluorescence Sensing[END_REF] . Moreover, the most important advantage of fluorescence is the versatility. There are plenty of different fluorescent materials, more are under development. All these features distinguish fluorescence from other methods and make it great importance in sensing and labelling.

As we discussed in §1.2, the fundamental issues for an ideal fluorescent material are brightness and stability. In the quest of the brightest and most stable fluorescence labels, different classes of materials are currently being developed and employed as fluorescent probes and they mainly include: organic dyes (Figure 1.6a), fluorescent proteins (Figure 1.6c)

and fluorescent nanomaterials (Figure 1.6b, d). The earlier classes of fluorescent labels such as organic dyes and fluorescent proteins, are still the most popularly used as fluorescent labels. This is because of their small size, high water solubility, commercial availability and the existence of standard protocols. However, there are several limitations for these labeling agents such as: broad spectrum profiles, photobleaching or poor photochemical stability [START_REF] Wang | Luminescent nanomaterials for biological labelling[END_REF] .

Nowadays, fluorescent materials with high brightness and high photostability are in great need for life science, at the same time, the developments of nanomaterial science bring a new class of labeling agent: fluorescent nanomaterials. Fluorescent nanomaterials have the great potential and ability to solve the drawbacks caused by "standard" fluorophores [START_REF] Wang | Luminescent nanomaterials for biological labelling[END_REF] . In this section I will concentrate on several important members of fluorescent labels: organic dyes, fluorescent proteins and fluorescent nanomaterials (e.g. semiconductor quantum dots, silica-based nanoparticles, and organic polymer nanoparticles). I will discuss their properties and evaluate their applicability for bioimaging and biosensing applications.

Organic dyes

Organic dyes are the most common probes used in fluorescence bioimaging. Their main advantages are their easy availability, low price and versatility. The amount of fluorescent synthetic organic dyes is quite large and keeps increasing, therefore, I will concentrate on several main classes that are frequently used.

Fluorescein is a green organic dye. It is widely used as a fluorescent label for biological applications. The chemical structure and spectra of the parent fluorescein dye is presented in Figure 1.7. Fluorescein has an absorption maximum at 494 nm and emission maximum of 521 nm in buffer at pH 7 41 . The pKa of fluorescein is 6.4 and its ionization equilibrium leads to a pH-dependent absorption and emission over the pH range of 5 to 9 [START_REF] Martin | The pH dependence of fluorescein fluorescence[END_REF] . Fluorescein and its derivatives are commonly used in confocal microscopy and flow cytometry applications.

However, fluorescein dyes have low photostability which is an obvious disadvantage for bioimaging application. There are many different fluorescein derivatives which are widely used as fluorescent labels, such as, fluorescein isothiocyanate (FITC) [START_REF] Twining | Fluorescein isothiocyanate-labeled casein assay for proteolytic enzymes[END_REF] , 6-FAM phosphoramidite [START_REF] Kutyavin | 3′-Minor groove binder-DNA probes increase sequence specificity at PCR extension temperatures[END_REF] , OregonGreen [START_REF] Rusinova | Alexa and Oregon Green dyes as fluorescence anisotropy probes for measuring proteinprotein and protein-nucleic acid interactions[END_REF] , Tokyo Green [START_REF] Mottram | The Pennsylvania Green Fluorophore: A Hybrid of Oregon Green and Tokyo Green for the Construction of Hydrophobic and pH-Insensitive Molecular Probes[END_REF] and SNAFL (Carboxyseminaphthofluorescein) [START_REF] Xu | A novel fiber-optic pH sensor incorporating carboxy SNAFL-2 and fluorescent wavelength-ratiometric detection[END_REF] . It has an absorption maximum at 507 nm and emission maximum of 529 nm in methanol 41 .

Its luminescence quantum yield is 0.90 [START_REF] Kubin | Fluorescence quantum yields of some rhodamine dyes[END_REF] . Rhodamine 123 seems to bind to the mitochondrion membranes and inhibit transport processes [START_REF] Johnson | Localization of mitochondria in living cells with rhodamine 123[END_REF] . Alexa Fluor dyes, a fluorescent dye family which is obtained by sulfonating aminocoumarin, rhodamine, or carbocyanine dyes [START_REF] Haugland | Handbook of Fluorescent Probes & Research Chemicals: Deutsche Mark Version[END_REF] . They are widely used as cell or tissue labels in fluorescence microscopy and cell biology. Alexa fluor dyes have a very wide excitation and emission spectra. They cover the whole visible spectrum and extend to the infrared range (Figure 1.9). Alexa Fluor dyes are normally more stable, brighter, and less pHsensitive than dyes like fluorescein, and rhodamine [START_REF] Panchuk-Voloshina | Alexa Dyes, a Series of New Fluorescent Dyes that Yield Exceptionally Bright, Photostable Conjugates[END_REF] . However, they are more expensive. BODIPY dyes (boron-dipyrromethene), is an important class of fluorescent dyes. The synthesis and spectroscopic properties have been recently summarized in a review by Loudet A et al. [START_REF] Loudet | BODIPY dyes and their derivatives: syntheses and spectroscopic properties[END_REF] . Normally, BODIPY molecules are uncharged [START_REF] Demchenko | Introduction to Fluorescence Sensing[END_REF] . The core structure of the BODIPY fluorophore is shown in Figure 1.10. BODIPY have shown attractive spectroscopic features such as tunable emission spectra from green to red. BODIPY dyes possesses uniquely small Stokes shift, high fluorescent quantum yield [START_REF] Karolin | Fluorescence and Absorption Spectroscopic Properties of Dipyrrometheneboron Difluoride (BODIPY) Derivatives in Liquids, Lipid Membranes, and Proteins[END_REF] . BODIPY dyes are highly soluble in many organic solvents. The combination of these characters makes BODIPY dyes an important tool in imaging applications. BODIPY dyes are considered to be very versatile [START_REF] Ulrich | The chemistry of fluorescent bodipy dyes: versatility unsurpassed[END_REF] .

Scientists can use BODIPY dyes to prepare BODIPY conjugated nucleotides 57 , proteins [START_REF] Schade | BODIPY-α-Casein, a pH-Independent Protein Substrate for Protease Assays Using Fluorescence Polarization[END_REF] , fluorescent enzyme substrates for life science [START_REF]The Molecular Probes® Handbook[END_REF] . Besides the dye families described above, there are also some fluorescent molecules used to target specific cell compartments or to label and distinguish dead and live cells.

Methylene blue is a blue fluorescent dye in its oxidized form, its maximum absorption is at 661nm, the maximum fluorescence emission is at 683nm (Figure 1.12). In water, methylene blue has a positive charge, and it will bind to DNA of bacteria which is negatively After focusing on molecular fluorophores, I will now make a short review about fluorescent proteins.

Fluorescent proteins

Fluorescent proteins (FP) are widely used in the life science, especially for probes and labelling in bioimaging applications [START_REF] Chudakov | Fluorescent Proteins and Their Applications in Imaging Living Cells and Tissues[END_REF] . The green fluorescent protein (GFP), a naturally fluorescent protein found from the jellyfish Aequorea Victoria, was the first in this family to isolated and characterized. The GFP from A. victoria has a major excitation peak at 395 nm and a minor one at 475 nm, its maximum emission peak is at 509 nm [START_REF] Tsien | The Green Fluorescent Protein[END_REF] In fluorescent proteins, the fluorophore lies within the folded protein, the mechanisms of fluorescent protein formation have been described by Miyawaki et al. [START_REF] Miyawaki | Mechanisms of protein fluorophore formation and engineering[END_REF] . The greatest advantage of fluorescent proteins is their heritability. Because of the ability to be co-synthesized and inherited inside the living cells, fluorescent proteins have become a very unique and important tool in living cell imaging [START_REF] Chalfie | GFP: Lighting up life[END_REF] . In recent years, GFP and its variants in different colors [START_REF] Shaner | Improved monomeric red, orange and yellow fluorescent proteins derived from Discosoma sp. red fluorescent protein[END_REF] have been widely used in the study of organization and function of living systems by optical techniques (Figure 1 .18). Fluorescent proteins encoded to a specific target in cells [START_REF] Mérola | Newly engineered cyan fluorescent proteins with enhanced performances for live cell FRET imaging[END_REF][START_REF] Erard | Minimum set of mutations needed to optimize cyan fluorescent proteins for live cell imaging[END_REF] , bacteria [START_REF] Andersen | New Unstable Variants of Green Fluorescent Protein for Studies of Transient Gene Expression in Bacteria[END_REF] , organisms [START_REF] Chudakov | Fluorescent proteins as a toolkit for in vivo imaging[END_REF] , animals [START_REF] Vintersten | Mouse in red: Red fluorescent protein expression in mouse ES cells, embryos, and adult animals[END_REF] allow for the study of localization, movement, function, mechanisms of this target. For example, fluorescent proteins can be used to label cell organelles, to investigate cell structure, or cell division, and to describe organelles functional process by microscopy [START_REF] Chudakov | Fluorescent proteins as a toolkit for in vivo imaging[END_REF]76,77 . Fluorescent proteins can also be used to label bacteria cells and to detect their movements, to study their transient gene expression process [START_REF] Andersen | New Unstable Variants of Green Fluorescent Protein for Studies of Transient Gene Expression in Bacteria[END_REF] and many other important characteristics [START_REF] Miroux | Over-production of Proteins inEscherichia coli: Mutant Hosts that Allow Synthesis of some Membrane Proteins and Globular Proteins at High Levels[END_REF][START_REF] Geoffroy | Use of Green Fluorescent Protein To Tag Lactic Acid Bacterium Strains under Development as Live Vaccine Vectors[END_REF][START_REF] Tombolini | Flow cytometric and microscopic analysis of GFP-tagged Pseudomonas fluorescens bacteria[END_REF] . However, the molecular mass of fluorescent proteins can reach 26-28 kDa, which is relatively large and may interfere with the function and localization of the labelled protein and the interaction with other molecules [START_REF] Valeur | Molecular Fluorescence[END_REF][START_REF] Demchenko | Introduction to Fluorescence Sensing[END_REF][START_REF] Wang | Luminescent nanomaterials for biological labelling[END_REF] . After describing fluorescent proteins as useful macromolecules for biochemists and biophysicists, I will now focus on larger objects such as fluorescent nanomaterials.

Fluorescent nanomaterials

The combination of fluorescence and nanomaterials has developed into an attractive and emerging interdisciplinary research field combining chemistry and photophysics.

Nowadays, fluorescent nanomaterials have attracted great interests due to their excellent optical properties including high photostability and high brightness. Furthermore, many developments have already been made to functionalize these nanomaterials to be more biocompatible. Additionally, most of the fluorescent nanomaterials possess tunable excitation and emission wavelength and can be used for multi-color imaging. The combination of these advanced features have made fluorescent nanomaterials to be an important and highly promising tool which can provide new roads of fluorescence imaging and sensing for in vitro and in vivo labeling in bacteria, cells, tissues, and whole organisms. In recent years, this approach has been widely used in various areas of biology and medicine, and environmental science. Fluorescent nanomaterials can be generally classified into two categories based on their chemical nature: inorganic fluorescent nanomaterials (e.g. quantum dots, [START_REF] Roy | Estimation of Minimum Doses for Optimized Quantum Dot Contrast-Enhanced Vascular Imaging In Vivo[END_REF][START_REF] Michael | Fluorescent Imaging Applications of Quantum Dot Probes[END_REF][START_REF] So | Self-illuminating quantum dot conjugates for in vivo imaging[END_REF] silica nanoparticles, [START_REF] Wu | Constructing NIR silica-cyanine hybrid nanocomposite for bioimaging in vivo: a breakthrough in photo-stability and bright fluorescence with large Stokes shift[END_REF][START_REF] He | Fluorescence Resonance Energy Transfer Mediated Large Stokes Shifting Near-Infrared Fluorescent Silica Nanoparticles for in Vivo Small-Animal Imaging[END_REF] carbon nanoparticles [START_REF] Jing Liang Li | A review of optical imaging and therapy using nanosized graphene and graphene oxide[END_REF][START_REF] Liu | Strong Two-Photon-Induced Fluorescence from Photostable, Biocompatible Nitrogen-Doped Graphene Quantum Dots for Cellular and Deep-Tissue Imaging[END_REF][START_REF] Demchenko | Novel fluorescent carbonic nanomaterials for sensing and imaging[END_REF] and metal nanoclusters [START_REF] Roy | Estimation of Minimum Doses for Optimized Quantum Dot Contrast-Enhanced Vascular Imaging In Vivo[END_REF][START_REF] Yu | Live Cell Surface Labeling with Fluorescent Ag Nanocluster Conjugates †[END_REF] ) and organic fluorescent nanomaterials (e.g. fluorescent conjugated polymer nanoparticles [START_REF] Fischer | Self-Assembled Fluorescent Organic Nanoparticles for Live-Cell Imaging[END_REF][START_REF] Howes | Phospholipid Encapsulated Semiconducting Polymer Nanoparticles: Their Use in Cell Imaging and Protein Attachment[END_REF][START_REF] Ding | Bright Far-Red/Near-Infrared Conjugated Polymer Nanoparticles for In Vivo Bioimaging[END_REF][START_REF] Pecher | Tailor-Made Conjugated Polymer Nanoparticles for Multicolor and Multiphoton Cell Imaging[END_REF] , dye-doped polymeric nanoparticles [START_REF] Thielbeer | Polymerizable Fluorescein Derivatives: Synthesis of Fluorescent Particles and Their Cellular Uptake[END_REF][START_REF] Kim | Single Step Reconstitution of Multifunctional High-Density Lipoprotein-Derived Nanomaterials Using Microfluidics[END_REF][START_REF] Reul | Near infrared labeling of PLGA for in vivo imaging of nanoparticles[END_REF][START_REF] Zhou | Fluorescent dextran-based nanogels: efficient imaging nanoprobes for adipose-derived stem cells[END_REF] ). I will focus on several representative fluorescent nanomaterials: quantum dots, silica nanomaterials, and organic polymer nanomaterials to discuss their properties and their biological applications.

The general steps to design functional fluorescent nanomaterials for biosensing and bioimaging are: 1) synthesis; 2) coating; 3) surface functionalization or bioconjugation; 4)

biosensing and bioimaging applications [START_REF] Yao | Biology, and Medicine of Fluorescent Nanomaterials and Related Systems: New Insights into Biosensing, Bioimaging, Genomics, Diagnostics, and Therapy[END_REF] Nowadays, QDs have become important nanoparticles for biological applications due to their unique optical properties that are unavailable from classical organic dyes and fluorescent proteins. QDs have high brightness, photostability, high quantum yield, and narrow emission. Their emission of light covers the visible and near infrared range 102,103 . In comparison with organic dyes, QDs have similar quantum yields but extinction coefficients that are 10-50 times greater and significantly more reduced photobleaching rates [START_REF] Yao | Biology, and Medicine of Fluorescent Nanomaterials and Related Systems: New Insights into Biosensing, Bioimaging, Genomics, Diagnostics, and Therapy[END_REF]104 .

When increasing the QDs size, their band gap decreases, this will shift their emission and imaging agents 100,107,115,116 . In comparison with traditional organic dyes, QDs are more photostable and brighter, however, the drawbacks such as irreproducibility, toxicity issues, blinking and complex fluorescence decays are to be considered when developing fluorescent labels with QDs [START_REF] Valeur | Molecular Fluorescence[END_REF] .

Thus chemists developed silica-based NPs which have less toxicity and higher fluorescence stability.

Silica-based nanoparticles

Another type of fluorescent nanomaterial which has been extensively developed for fluorescent labelling and bioimaging is silica nanoparticles. Generally speaking, silica provides a stable vehicle to protect the encapsulated fluorophores from external perturbations 123 . The silica nanoparticles matrix is capable of encapsulating a variety of fluorescent agents such as inorganic dyes [124][125][126] , organic dyes 127-129 (Figure 1.25), QDs and metal NPs without affecting their spectral features [START_REF] Yao | Biology, and Medicine of Fluorescent Nanomaterials and Related Systems: New Insights into Biosensing, Bioimaging, Genomics, Diagnostics, and Therapy[END_REF]124,130 . Recently, multi-shell silica nanoparticles have also been developed for multi-functional purpose in drug delivery and fluorescent labeling applications. For example, the multi-shell architecture displayed in Figure 1.27 is formed with a gold nano shell for near infrared plasmon imaging and a silica outer shell for drug delivery. By varying the chemical properties of each layer, this kind of particle can be used for different tasks such as drug delivery, bioimaging and sensing 141 . Compared with conventional organic dyes, silica nanoparticles are more hydrophilic, biocompatible and more stable. However, silica particles also have some drawbacks, such as particle aggregation 142 and organic dyes spill-over from particles 36 that may limit their applications.

Although inorganic nanomaterials have been intensively developed for fluorescent imaging in vitro, the potential bioincompatibility of the inorganic nanomaterials have hindered the development of their practical applications for imaging in vivo [143][144][145] . Nowadays, the U.S. Food and Drug Administration(FDA) has only approved one inorganic nanomaterial:

the nonporous silica nanoparticle for the phase I clinical trial 146 . In contrary, organic fluorescent nanomaterial imaging agents are obtained from organic molecules that are less toxic. Furthermore, in order to develop organic materials, there are already many FDAapproved biocompatible molecules, such as the fluorescent dye indocyanine green (ICG) 147 and the biodegradable polymers poly(methyl methacrylate) (PMMA), poly(lactic acid) (PLA), poly(lactic-co-glycolic acid) (PLGA) derivatives and poly(ethylene glycol) (PEG) 148 . In the next section, I will discuss the organic fluorescent nanomaterials.

Organic polymer nanoparticles (OPNs)

It is well known that the major problem with traditional organic dyes is their poor photostability, this major drawback results in low sensitivity for biological labeling and sensing studies. Fluorescent nanomaterials are brighter and more photostable compared with classical organic dyes. This is because every nanoparticle can carry a large number of fluorescent species which are insulated by the particle matrix. Indeed this matrix can protect enfolded fluorophores from complex environment. Moreover, the large surface to volume ratio of nanoparticles offers a suitable platform for surface modification and Organic polymer nanoparticles are known as highly versatile nanostructured materials.

As we discussed above, the advantages of organic polymer nanoparticles includes high brightness, high photostability, high quantum yield, low toxicity, and rich surface modification. In order to prepare organic polymer nanoparticles, there are several methods available: macro-/micro-/miniemulsion, precipitation, self-assembly, direct polymerization.

Each of them will be described below.

Methods of synthesize organic polymer nanoparticles

Landfester and co-workers have synthesized organic polymer nanoparticles by the miniemulsion method 152,153 (Figure 1.29). To prepare organic polymer nanoparticles, the polymer is dissolved in an organic solvent and then the solution is injected into an aqueous solution of an appropriate surfactant 154 . The mixture is then rapidly stirred by ultrasonicating to generate stable miniemulsion which contains small droplets of the polymers. Finally, the organic solvent is evaporated and leaving a stable dispersion of polymer nanoparticles in water. Depending on the polymer concentration, the size of this kind of particle can vary from 30 to 500 nm 155 . During precipitation, hydrophobic conjugated polymers are dissolved in a good solvent such as THF, and then the mixture is poured into a poor solvent such as water which is miscible with the good solvent (Figure 1.30). The main force for the formation of the nanoparticles is the hydrophobic effect. When the solution of polymer in good solvent is added into the water, polymer chains tend to avoid facing with water and fold into spherical shapes in order to achieve minimum exposure to water. The solution is also stirred powerfully with a sonicator to help the formation of nanoparticles 155 . The particle diameter is dependent on the starting concentration of the conjugated polymers 156,157 . Higher polymer starting concentration will result in larger particle diameter. In direct polymerization, monomers and organic dyes are dissolved in the organic solvent, and then the mixture is added into the aqueous solution with an emulsifier. With ultrasonification, stable and small oil droplets will be formed inside the mixture solution.

After adding initiators into the emulsion, monomers in oil droplets will start the polymerization to obtain organic nanoparticles dispersion (Figure 1.32) 158,159 .

Fluorescent labelling and sensing applications

After a brief description of the methodologies of preparation of organic polymer nanoparticles, I will describe the main applications of these ONPs for fluorescent labelling and fluorescent sensing.

As I have described before, due to their biocompatible natures, organic polymer nanoparticles have relatively low cytotoxicity compared with inorganic nanoparticles. Small organic polymer nanoparticles can be internalized by living cells through the process of endocytosis without extra modification of the particle surface or addtional technologies.

Various fluorescent organic nanoparticles have been used to label cells including fluorescent conjugated polymer nanoparticles [START_REF] Fischer | Self-Assembled Fluorescent Organic Nanoparticles for Live-Cell Imaging[END_REF][START_REF] Howes | Phospholipid Encapsulated Semiconducting Polymer Nanoparticles: Their Use in Cell Imaging and Protein Attachment[END_REF][START_REF] Ding | Bright Far-Red/Near-Infrared Conjugated Polymer Nanoparticles for In Vivo Bioimaging[END_REF][START_REF] Pecher | Tailor-Made Conjugated Polymer Nanoparticles for Multicolor and Multiphoton Cell Imaging[END_REF] , self-fluorescent biodegradable nanoparticles 161 , dyedoped polymeric nanoparticles [START_REF] Thielbeer | Polymerizable Fluorescein Derivatives: Synthesis of Fluorescent Particles and Their Cellular Uptake[END_REF][START_REF] Kim | Single Step Reconstitution of Multifunctional High-Density Lipoprotein-Derived Nanomaterials Using Microfluidics[END_REF][START_REF] Reul | Near infrared labeling of PLGA for in vivo imaging of nanoparticles[END_REF][START_REF] Zhou | Fluorescent dextran-based nanogels: efficient imaging nanoprobes for adipose-derived stem cells[END_REF]J.H. et al. 162 Organic polymer nanoparticles can also be used to coat the bacterial surface to form bacteria-nanoparticle microparticles to perform multiplex biological imaging and analysis.

Wang and co-workers 163 prepared four different colored conjugated polymer nanoparticles (P1 blue, P2 green, P3 yellow and P4 red) with diameters from 50 to 100 nm. By electrostatic and hydrophobic interactions, E. coli bacteria were coated with particles. These fluorescent organic nanoparticles were bounded to the outer membrane of E.coli. Under excitation from a single wavelength, multi-step intermolecular FRET processes between these particles occurred. By monitoring the proportions of P1, P2, P4 on E. coli bacteria, different colored bacteria-nanoparticle microparticles can be obtained (Figure 1.34). Because of their nontoxic properties, organic polymer particles can be used not only for cell imaging but also for drug delivery 164,165 Besides electrostatic interactions between organic polymer particles and bacteria, bioconjugation of the particles surface with various biological functional groups such as avidin, biotin 168 and antibodies 169,170 allows them to target the bacterial surface for various biological applications. The surface of polystyrene (PS) beads can be coated by antibody 171- 173 or streptavidin 174 . Rashid Bashir and his group 174 have used streptavidin-coated nanoparticles which can bind with a biotinylated-antibody to target bacteria in order to form a bacteria cargo for tracking and drug delivery application as displayed in Organic polymer nanoparticles can also be used for bacterial detection and monitoring bacterial growth. The growth of bacteria is often associated with a pH decrease of the growth medium due to a release of acidic metabolites such as acetic acid, lactic acid and CO 2.

Different kinds of organic nanoparticles pH sensors have therefore been used to measure the growth of bacteria based on this principle 175,176 . Bhawana Thakur et al. 175 demonstrated that a polyaniline nanoparticle based colorimetric pH sensor could monitor bacterial growth.

Polyaniline is highly sensitive to the presence of protons and it will change from blue to green visible color. In this work, polyaniline nanoparticles were synthesized and The core-shell structure fluorescent nanoparticles were synthesized and characterized by Chloé Grazon [START_REF] Grazon | Élaboration de nanoparticules fluorescentes à base de BODIPY par polymérisation RAFT en miniémulsion : synthèse, caractérisation et fonctionnalisation de surface[END_REF] , a former PhD student in PPSM (CNRS UMR8531) laboratory. One particle is formed with around 1750 amphiphilic self-assembled polymer chains bearing about one to two BODIPY per chain. Thus about 3000 BODIPY fluorophores are inside the core, this resulted in very high brightness. These FNPs are also stable and photobleach slowly due to the protection of fluorophores in the hydrophobic core. Thanks to the poly-acrylic acid groups all around the particle, it can be functionalized easily with biomolecules such as biotin and antibodies.

These FNPs are an ideal tool for biolabeling and bioimaging applications in life science due to their high brightness, high photostability and rich surface functionalization ability. In FNP-bioconjugation). The third strategy is that fluorescent nanoparticles can be used as pHbiosensor to detect the growth of bacteria for biosensing application (Figure 1.40E: FANPs).

In this manuscript I described the investigation of the biocompatibility of FNPs on E.coli bacteria. I will describe a methodology to insert the described FNPs ( I then performed bioconjugation of biotin to FNPs (Figure 1.40B). Biotin conjugated FNPs can target bacteria surface membrane. By using a streptavidin-biotin link, I can obtain a "sandwich" to build a bridge between particles, specific antibodies and bacteria. SPR, fluorescence images and SEM images have been used to demonstrate the interaction of biotin conjugated FNPs with E.coli bacteria. An antibody specific to the bacterial outer membrane can also be conjugated on the particle surface, I will describe how we can use these antibody coated FNPs to target bacteria (see Chapter 3).

We also have designed green fluorescent polymer chains (GFPCs, After a general introduction, I will describe in details the interation of these fluorescent nanomaterials with E.coli bacteria and the bioimaging and biosensing applications of them on bacteria in following chapters. Now, I will begin with fluorescent nanoparticles (FNPs) in chapter 2, I will show you three different fluorescent nanoparticles and the methology to insert them inside bacteria for internal bioimaging of E.coli.

Chapter 2 Fluorescent nanoparticles (FNPs) for internal bioimaging of E.coli bacteria

In this chapter, I will present three different fluorescent nanoparticles: one negatively charged (FNP -) and two positively charged fluorescent nanoparticles (FNP + and FNP-PEG + ). I will discuss their photophysical characterizations and use for internal bioimaging of E.coli bacteria.

Synthesis of fluorescent nanoparticles (FNPs)

The synthesis of the fluorescent nanoparticles presented here has been optimized by Dr. Chloé Grazon 180 . The one-pot synthesis is based on a reversible addition-fragmentation chain-transfer (RAFT) polymerization in miniemulsion conditions using a surfactant-free onepot phase inversion process (Scheme 2.1) 180 .

Briefly, the particles are obtained from a hydrophilic chain PEO-b-PAA (poly(ethylene oxide)-block-poly(acrylic acid)) macromolecular RAFT agent (experimental details in Chapter 7, §7.2.1). These hydrophilic chains were first extended with a hydrophobic block by a bulk polymerization of styrene (S) and BODIPY methacrylate-1 (BDPMA1). Then, the polymerization was stopped after 70 minutes. Basic water (pH 12.5) was added to the reaction mixture at room temperature and ultra-sound treatment in an ice bath allowed the formation of nano sized monomer droplets (amphiphilic copolymer PEO-b-PAA-b-P(S-co-BDPMA1)) were obtained. Finally, the RAFT polymerization was reinitiated by simple degasing with argon and heating to 80°C. After re-polymerization, a core-shell structure negatively charged nanoparticle was obtained with poly(styrene-co-BDPMA1) in the core and poly(ethylene oxide)-block-poly(acrylic acid) in the shell. The particle is negatively charged in water because the carboxylic acid group (-COOH, pKa around 4) will dissociate into H + cations and (-COO -) anions in water (pH 6.8-7.0). The obtained FNP -are well-defined in size with a mean hydrodynamic diameter of 65 nm (Table 2.1).

Scheme 2.1 Synthetic path way for fluorescent nanoparticles (FNP -, FNP + , FNP-PEG + )(adapted from 180 ).

In order to obtain positively charged particles, the acrylic acid sites on FNPs were reacted with ethylenediamine or poly(ethylene glycol) diamine by a coupling reaction involving activation of the carboxylic acid with N-(3-Dimethylaminopropyl)-N'ethylcarbodiimide hydrochloride (EDC-HCl) (Scheme 2.1). These particles are positively charged in water because the amine group (NR 3 , pKa around 9) will dissociate into N + cations and (R 3-) anions in water (pH 6.8-7.0). Two different positive fluorescent nanoparticles were obtained: FNP + and FNP-PEG + with very small difference on the shell where FNP-PEG + has slightly longer side chains than FNP + as shown in Scheme 2.1.

Characterizations of fluorescent nanoparticles (FNPs)

Fluorescence and absorption spectroscopy analysis of FNPs are displayed on Figure 2.1

and their main spectroscopic data are given in Table 2.1. The BDPMA1 monomer in toluene shows the expected spectra for BODIPY fluorophores [START_REF] Loudet | BODIPY dyes and their derivatives: syntheses and spectroscopic properties[END_REF] with an intense band in the visible region located at 528 nm (corresponding to  transition) and a vibrational shoulder at higher energy. A second, less intense band is located in the UV region at around 380 nm 180 .

The maximum of fluorescence emission is found at 540 nm, exhibiting a Stokes shift of 421 cm -1 (12 nm).

The FNP -exhibits very similar absorption and fluorescence spectra with a maximum absorption at 529 nm and a maximum fluorescence emission at 543 nm. The Stokes shift of the nanoparticles (487 cm -1 , 14nm) is slightly higher than the monomer. Similar slight red shift of fluorescence emission has previously been observed from free monomer to monomer immobilized in a rigid environment 181 . One can also notice that the FNP - absorption spectra exhibit a more intense vibrational shoulder which might result from the confinement of BODIPY in a rigid polymer matrix (Figure 2.1A) 181,182 . The excitation spectra for FNP -and BDPMA1 monomer have the same aspect and superimpose to their respective absorption spectra (Figure 2.1B), showing that the absorbing and emitting species are identical. The fluorescence quantum yield of FNP -is 36% while that of BODIPY in toluene is 70%. The decrease may be due to quenching between closely associated BODIPY monomers in the polystyrene matrix. However since there are about 2975 BODIPY molecules 27 inside one particle, the brightness of the particle is about 7.8x10 7 M -1 cm -1 , which is 1500 times higher than the one of BODIPY monomer alone. The very high brightness of FNP -shall be a good advantage for bioimaging applications (see Chapter 1 §1.2).

A B Φ F /% toluene

Toxicity assessments of FNPs on E.coli

Having synthesized and characterized the FNPs, I next assessed the toxic effect of FNPs on E.coli. In order to do this, the growth rates of bacteria in the presence of FNPs were compared with the growth rate of E.coli bacteria alone. The growth rate was measured by using a 96-well plate reader. The FNPs concentration used was 6.6x10 -3 µM (4x10 12 particles/mL).

Here, E.coli bacteria with FNP -, FNP + or FNP-PEG + were incubated in the 96-well plate reader at 37°C in the dark overnight. A control with only bacteria was also prepared. Every nine minutes the growth rate of the cells was monitored by measuring the optical density at 600nm (Figure 2.4). For over ten hours incubation, the growth curves of bacteria with FNPs were practically the same as the growth curve of bacteria alone both in LB (rich) and M9

(poor) media. This means that, at this concentration, FNPs have no toxicity effect on bacterial growth in a long time scale, independently of the charge of the FNPs.

FNPs for internal bioimaging of E.coli

Chemical treatments of E.coli with FNPs

After having shown that FNPs have no toxicity effect on E.coli bacterial growths, the FNPs were tested for internal imaging application with E.coli. The Escherichia coli cytoplasmic membrane is composed of two phospholipid bilayers with a peptidoglycan layer sandwiched between them and has the function of acting as a permeability barrier for most molecules. There are two means of transportation of molecules or biochemical agents into the cell: active transport or passive transport. Passive transport is a movement of molecules across the bacterial membrane. It is driven by the increase of entropy of the system. Active transport is also a movement of molecules (such as ions, glucose and amino acids…) across bacterial membrane in the direction from lower concentration to higher concentration.

Unlike passive transport it requires an input of cellular energy. Particles under 5nm can be transported into bacterial cell by active transport 183 . As a matter of fact, the size of FNPs (60-80nm) is too big to enter inside E.coli and we need to find another way to insert them. One way to remove the limitation of membrane barrier is to partially destroy it 184 . Historically, a variety of chemicals were found to penetrate E.coli. For example, the cell membrane may be gently lysed and disorganized by chemical treatment with di(ethylene glycol) (DEG) 185 . Thus, in the following experiments, E.coli bacteria are permeabilized by a brief exposure to DEG before incubation with FNPs.

An initial E.coli bacterial concentration of 5x10 7 CFU/mL (OD=0. After incubating E.coli with FNP -and DEG, the percentage of total fluorescent cells increased when increasing the FNP -concentration and it reaches a plateau when the FNP -is about 3.9x10 -2 µM. The percentage of total fluorescent cells after DEG treatment could reach 80%. I choose the FNP -concentration of 2.6x10 -2 µM as a suitable concentration to continue the chemical treatment experiments. Since the chemical treatment was not efficient to insert FNPs inside E.coli bacteria, I turned to another method: electroporation.

Electroporation of E.coli bacteria with FNPs

Characterization of electroporation by flow cytometry

Another method to partially destroy the membrane barrier in order to insert the FNPs into the E.coli is electroporation. Electroporation consists in the application of an electric pulse on the bacterial cell, pores will form on the cell membrane, and the fluorescent particles may enter inside the membrane through these holes. During incubation, cells can repair their membranes while the particles stay inside the cell. Electrocompetent cells are needed for this process; competence is the ability of a cell to take up extracellular elements or materials from its environment. Electrocompetent cells are prepared by a chemical treatment (see experimental part for details in Chapter 7, §7.5.2). 50µL electrocompetent cells (8x10 7 bacteria /µL) were mixed with different volumes of FNP -, FNP + , FNP-PEG + to reach different final concentrations (2.7x10 -4 -5.2x10 -2 µM). This mixture was placed on ice for 10 minutes before electroporation at the specified voltage (15kV/cm). Immediately after electroporation, the cells were then incubated at 37°C for one hour in rich growth medium to repair their membranes.

After incubation, cells were centrifuged to remove unattached nanoparticles and fixed with 4% formaldehyde. Then the cells were re-suspended in PBS (1mL) prior to flow cytometry measurement and bioimaging.

Fluorescence bioimaging of E.coli interacting with FNPs after electroporation

E.coli's interaction with FNP -, FNP + , FNP-PEG + was assessed by fluorescence microscopy.

Phase contrast images and fluorescence images were taken. E.coli cells interact also with positively charged FNP + and FNP-PEG + (Figure 2.12). As it was observed with the FNP -, most of the cells are associated with particles, and several FNPs on one E.coli bacterium are often observed. As mentioned previously we cannot say whether FNPs are on the membrane or inside the bacteria. From the images and as already observed by flow cytometry, we found that the probability of the interaction of E.coli with FNPs is not dependent on the particle charge. After observation of FNPs interaction with E.coli bacteria, we found that the FNPs can be associated with the cell after electroporation. However, only 8% of cells are fluorescent and alive. Furthermore, it is difficult to determine whether the particles are inside the bacteria from the microscopy images, since they could also be on the surface of bacteria. So, in order to discriminate between these two hypotheses, another dye, methylene blue, was introduced in the experiments.

Emission quenching of FNP -with Methylene Blue (MB)

In order to determine whether after electroporation the FNPs were inside or outside E.coli, methylene blue was used. Methylene blue (MB-see Chapter 1 §1.3.1) is a cationic blue biocompatible dye, it will bond to the DNA of bacteria which is negatively charged.

Methylene blue is reduced in viable cells leaving them unstained, dead cells are unable to reduce the oxidized methylene blue and the cells are stained blue. However, the labeling of bacteria by methylene blue is requires a longer time (around 10 minutes) 186 compared with the quenching process and a particular protocol is needed for the staining process 187,188 , so it is possible to use the methylene blue to quench the particles outside the bacteria. I will first demonstrate that MB can interact with FNPs (without bacteria) and quench their emission.

Here, I take FNP -as an example to continue the following experiments since the probability of the interaction of E.coli with FNPs is not dependent on the particle charge. The results of the fluorescence and absorption spectroscopy analysis of FNP -and methylene blue are displayed on Figure 2.13A. The overlap between the emission spectrum of FNP -and the absorption spectrum of MB indicates a possible quenching process of FNP -by MB. Titration experiments were conducted to characterize the quenching process of FNP -by MB. A FNP - solution was placed inside a cuvette and the methylene blue solution was added gradually.

Emission spectra were taken after each addition until the emission of FNP -stopped decreasing.

The titration experiment of FNP -with methylene blue is displayed on In order to understand the mechanism of the quenching process, the data was plotted according to Stern-Volmer equation shown in

Fluorescence bioimaging of E.coli with FNP -after quenching with MB

After characterization of the quenching process of FNP -, methylene blue was used in the bioimaging experiments of E.coli electroporated with FNP -. E.coli's interaction with FNP - after the addition of methylene blue was measured by epifluorescence microscopy. So, if there is a bacterium with particle inside and outside (Figure 2.15), after the addition of methylene blue, the particles outside the bacteria will be quenched and cannot be detected under the microscope, however the particles inside the bacteria will be protected by the membrane and cannot be quenched by the methylene blue due to that MB interaction time with bacteria is too short to go through the cell membrane. As a result, they will remain fluorescent and detected under microscope. After adding methylene blue, indeed, the FNP -outside the cells were quenched while some bright spots remain. This means that there were FNP -inside the cells. They were protected by the cell membrane and thus remained fluorescent under microscope (Figure

2.15B).

This means that FNP -can successfully enter inside E.coli bacteria after electroporation. However, when I tried to incubate bacteria with FNP -inside under the microscope, they did not grow. This is probably because, after electroporation, most of the cells were dead as determined by flow cytometry experiments, where only 8% cells were found alive and fluorescent. It is then difficult to identify them and watch them growing under the microscope.

SEM images of E.coli interacting with FNP -

Scanning electron microscope (SEM) was used in order to get more detailed information on E.coli interacting with FNP -after electroporation. A scanning electron microscope (SEM) is a type of electron microscope that produces images of a sample by scanning it with a focused beam of electrons. The resolution of SEM images can reach 1nm.

For SEM, a specimen is normally required to be completely dry, since the specimen chamber is under high vacuum. But living cells and tissues are soft and hydrated, so cells need to be fixed to be transformed into a solid state to preserve and stabilize their structure.

Fixation is usually performed by incubation in a solution of a buffered chemical fixative (with a fixing solution). In our case we used glutaraldehyde which kills cells quickly by crosslinking their proteins. The fixed tissues are then dehydrated. However air-drying in SEM is not suitable because air-drying causes collapse and shrinkage. For example, for fresh porcinis, when water is evaporated, high surface tension of water leads to a collapse of structures and air dried porcini are shrinking (Figure 2 After substitution of ethanol, it was exchanged with a transitional liquid carbon dioxide by critical point drying. Generally speaking, ethanol was used to replace water inside cells, and then ethanol was exchanged by high pressure liquid carbon dioxide. The liquid carbon dioxide is then heated until its temperature goes beyond the critical point, at which time the pressure can be gradually released, allowing the gas to escape and leaving a dried bacteria. A thin heavy metal platinum layer is applied to the specimen surface by coating in order to increase contrast by localization of the signal to the surface of the specimen.

As described in the beginning of electroporation experiments of bacteria ( §2.

Conclusion

In this chapter, I presented three different fluorescent nanoparticles: negatively charged fluorescent nanoparticle (FNP -), positively charged fluorescent nanoparticles (FNP + , FNP-PEG + ) and their bioimaging on E.coli bacteria.

FNPs have been shown to be nontoxic on bacterial growth in a long time scale independently of the charge and of the growth medium. Electroporation was used to insert particles inside bacteria. After electroporation, the percentage of total fluorescent cells could reach 90%, but because of the electroporation and the addition of FNPs, the percentage of live florescent cells decreased to about 8%.

Bioimaging experiments indicated that E.coli bacteria interacted with both negative and positive fluorescent nanoparticles, most of the cells are associated with particles. In order to make sure that FNPs were really inside the bacteria, methylene blue was introduced to quench the fluorescence of particles outside the bacteria. FNP -inside the cells were protected by the cell membrane and remained fluorescent after the addition of methylene blue while the particles outside the membrane could be quenched. This indicates that FNP - can successfully enter inside bacteria after electroporation.

SEM images of E.coli electroporated with FNP -have also shown that particles could be easily found on or under the membrane of bacteria, but bacteria are too much stressed and most of them were dead.

Only 8% cells are alive and fluorescent after electroporation which makes it difficult to identify them and watch them grow under the microscope. So, new strategies are needed to apply the particles to bioimaging of E.coli. One way is to try to decrease the size of the nanomaterials and synthesize smaller nano-objects for internal bioimaging of E.coli. Another way is to use the particles for external bioimaging of E.coli, for instance, we can try to link particles to the surface of bacteria which will be described in detail in next chapter. Finally, the labeling of dead bacteria by the FNP -could be used to measure the toxicity of potential antibiotics in a very sensitive manner due to the high brightness of the particles.

Additional images

Chapter 3 Fluorescent nanoparticles (FNPs) for external bioimaging of E.coli bacteria

In the previous chapter, I have shown that particles can be successfully inserted inside bacteria by electroporation, however, only very few bacteria survived after this treatment.

So, new strategies were needed to develop particles applied to fluorescent bioimaging of E.coli. If it is difficult to use them for internal labeling, could they be useful for external labeling of E.coli bacterial surface? The answer is yes. In this chapter, I will show how I modified the particles to link them to E.coli surface through the elaboration of a "sandwich system". Full characterization of the targeting process will be presented. Moreover the first step toward a microfluidic device designed to immobilize E.coli on a surface for further bioimaging will be shown. I also prepared a "nanoparticle-antibody" system to bind to the surface of E.coli bacteria. Preliminary imaging results of "nanoparticle-antibody" system will be presented.

Design of "Sandwich" for targeting bacterial surface

Conjugation of biotin on FNPs

After studying fluorescent nanoparticles for internal bioimaging application of E.coli bacteria, FNP -were then modified for bioimaging of bacterial surface. In order to target E.coli's outer membrane with FNP -, a "sandwich system" was created (Scheme 3.1). E.coli can be easily targeted with an antibody conjugated with biotin. The binding of biotin to streptavidin is one of the strongest non-covalent interactions known in nature. If we can conjugate biotin to nanoparticles, then with a streptavidin, we can easily use the particle based "sandwich system" to target the bacteria E.coli's outer membrane. After conjugation of biotin on FNP -, FNP --biotin was then used to target bacterial membrane by building the "sandwich system" (see experimental details in §3.1.2). Briefly, E.coli bacteria were firstly labelled by a primary antibody which was already conjugated with biotin. Then, the primary antibody conjugated with biotin was bonded to streptavidin. Since streptavidin has four identical peptide chains which can each bind to a biotin, the FNP --biotin was added to bond the free streptavidin sites. Finally, by using a streptavidin-biotin link, I built a bridge between particles, antibodies and bacteria (Scheme 3.3).

Spectroscopy analysis

In order to characterize the formation of the sandwich, I divided our study into four different parts: FNP -, FNP --biotin, FNP --biotin-streptavidin, FNP --biotin-streptavidin-antibody.

I analyzed each step of the sandwich formation, and did a comparative study of the properties of the subunits.

Fluorescence and absorption spectroscopy analysis of FNP -, FNP --biotin, FNP --biotinstreptavidin, FNP --biotin-streptavidin-antibody are displayed on Figure 3.1 and their main spectroscopic data are given in Table 3.1. The FNP -exhibits a maximum absorption at 529nm and a maximum fluorescence emission at 543nm. After conjugation of biotin, bonding with streptavidin or bonding with antibody didn't change the position of the maximum of absorption and emission. The only difference one can notice is that, FNP --biotin-streptavidin and FNP --Biotin-streptavidin-antibody exhibit a more intense band located in the UV region at around 380nm compared with FNP -and FNP --biotin which may come from the light scattered by the larger sized "sandwich" compared to the starting FNP -. From literature, we can find that the sizes of biotin, streptavidin and antibody are about 1nm 190 , 5nm 191,192 and 10 nm 193,194 respectively. Since the size of FNP -is about 65nm, theoretically, the formation of FNP --biotin-streptavidin and FNP --biotin-streptavidin-antibody complexes should give objects with a size of 77nm and 97nm respectively, much larger than FNP -and FNP --biotin (67nm) resulting in an increase of scattered light. In PBS the fluorescence quantum yield of FNP -is 36% and there is practically no difference for FNP --biotin (35%), FNP --biotin-streptavidin (34%) and FNP --biotin-streptavidinantibody (30%). Thus formation of the "sandwich" does not markedly alter the photophysical properties of FNP -. From the DLS measurements (Figure 3.2), one can notice that there is very little difference between the theoretical diameter and the hydrodynamic diameter (Table 3.

1).

This is likely to prove that the addition of each element is effective. There is only a slight deviation after the addition of antibody. This is probably because an antibody is relatively large (150 kDa 194 ) and it is difficult for them to cover the whole surface of a particle. Thus it is likely that only a few antibodies bond to the surface of particle resulting in a smaller size than the theoretical one for the FNP --biotin-streptavidin-antibody complex.

Surface Plasmon Resonance

Surface Plasmon Resonance (SPR) experiments (cooperation with Dr. Claude Nogues † )

were performed to measure the affinity of streptavidin and the FNP --biotin. SPR is a method that measures the interaction between a ligand and a receptor bound to a metallic surface.

The detection is based on the refractive index variation during the bio-molecular interaction.

Streptavidin solutions at different concentrations (10 µM and 1 µM in phosphate buffer (0.4 M) at pH 7.4) were spotted on a prism surface freshly covered with gold (Figure the SPRi apparatus and a 1 nM FNP -solution was flowed. Different images before and after the injection were taken. After the washing, a 1 nM FNP --biotin solution was flowed across the SPRi surface containing the spotted streptavidin and different images were taken before and after the injection. Therefore, the non-specific interaction of the FNP -with streptavidin was not strong enough and particles have been washed away. After the washing process, 1 nM FNP --biotin solution was flowed across the SPRi surfaces containing the spotted streptavidin. The kinetic curve of FNP --Biotin and streptavidin interaction confirms this observation (Figure 3.5B). The curve increased dramatically at the beginning and reached quickly a maximum proving the strong non-covalent interaction between particles and streptavidin.

Moreover, there is a higher relative reflectivity intensity for the spots at 10 µM streptavidin than the one at 1 µM which indicates that increasing the streptavidin concentration increases the non-covalent bonding between particles and streptavidin. Immediately after injection of FNP --biotin, the surface was continuously rinsed with PBS buffer. However the background signal remains while the relative reflectivity intensity in the streptavidin containing spots remained high unlike the ones flowed with FNP -. Similarly on the kinetic curve, the relative reflectivity intensity remained constant and didn't decrease at all even during washing (Figure 3.5B). Therefore, the non-covalent interaction between FNP --biotin and streptavidin is much stronger compared to the non-specific interaction of the FNP -and streptavidin. Moreover, the SPR experiments also proved the successful conjugation of biotin on FNP -.

Secondary fluorescent antibody

The efficiency of primary antibody to target E.coli bacteria was measured by

Fluorescence bioimaging of "sandwich" labelling bacterial surface

After characterization of each part of the "sandwich" system by spectroscopy, DLS, SPR and secondary antibody labeling, I demonstrated each part of the "sandwich system" are functionable and the "sandwich system" were successfully obtained. I applied it to label the bacterial membrane. Bioimaging of the particle based sandwich system targeting bacterial membrane were measured by fluorescence microscopy. However, even if we proved the formation of the "sandwich" system, it was very difficult to observe many cells targeted by particles. Moreover it was difficult to tell if the particles were really linked to the surface of bacteria.

In order to confirm the interaction of the sandwich with the bacteria, SEM images were also recorded.

SEM images of "sandwich" labelling bacterial surface

Scanning electron microscope (SEM) was used in order to get more detailed information on the sandwich system at the surface of E.coli. The cells were prepared like Without adding streptavidin, the sandwich is not built, and particles are hardly found on the membrane of bacteria (Figure 3.9A, B). When adding streptavidin, the sandwich is built and, we can find particles bound to the surface membrane of the bacteria which indicates the successful formation of the sandwich system (Figure 3.9C, D).

Sandwich complex in a microfluidic device

In order to improve and also to develop a new device to target bacteria, microfluidic system was introduced in our experiments. From previous experiments, I often observed a strong affinity of FNP -for glass surface: particles tended to attach to microscope slides. They proved out to be difficult to wash away after bonding. This unique property gave us the idea to build a sandwich system on a microscope slide to "catch" bacteria and target bacterial membrane more efficiently than in solution.

In order to measure the affinity of FNP --biotin particle to glass and to determine which suitable FNP --biotin amount is needed, a 6 channel μ-Slide (µ-Slide VI 0. After optimizing and obtaining a suitable FNP --biotin particle surface, streptavidin solution (4nM) was injected to the channel in order to bind the FNP --biotin particles. After gently washing away the unattached streptavidin, primary antibody solution (88nM) was injected to the channel to bind the streptavidin and build the "sandwich system". Finally, 30 µL bacterial solution with an initial concentration of 5x 10 7 CFU/mL (OD=0.5) was injected inside the channel and left for about one hour at room temperature. The whole process is depicted in Figure 3.12.

Figure 3.12 Process to form the "sandwich system" on a microscope slide.

It is not possible to detect bonding of streptavidin and antibody to FNP --biotin by microscopy. However, after one hour incubation, bacteria filled the channel as shown in Moreover every bacterium is linked to particles indicating that the "sandwich system" on microscope slide can "catch" bacteria and target bacterial membrane effectively.

Design of "Nanoparticle-Antibody" for targeting bacterial surface

In the "sandwich system", it takes three agents to link the FNP -to antibody which can target bacteria (FNP --biotin-streptavidin-biotin-antibody). Even if it is proved that "sandwich system" was successfully obtained, the targeting efficiency on bacteria is not very high. This may come from the several formation steps of "sandwich". Is it possible to link FNP -directly on primary antibody? The answer is yes. On the surface of antibody , there are a lot of amino acid residues 198 . Since the FNP -surface is covered with acrylic acids groups, we can conjugate directly antibody on the particles' surface by coupling reaction.

As I described in Chapter 2, FNP -is a core shell structure nanoparticle with poly(styrene-co-BDPMA1) in the core and poly(ethylene oxide)-block-poly(acrylic acid) (PEOb-PAA) in the shell (Figure 3.14A). The poly(acrylic acid) block used in the particle is designed for post-functionalization. In the "sandwich system", I introduced biotin conjugated to these acrylic acids units. However, from the structure displayed in The chemical bonding of the antibody to the nanoparticle was possible starting from amino acid residues on antibody that can react with the carboxylic acids present on the surface of FNPB. It was thus similar to the reaction used to transform FNP -into FNP + presented in Chapter 2 ( §2.1). In a typical experiment, the total number of acrylic acid units present on the surface of the branch chains fluorescent nanoparticles (FNPB) was taken as 1 equiv. Anti-E.coli antibody (4 equiv.) and EDC.HCl (4 equiv.) were added to react with the acrylic acids on the surface of the FNP -and get maximum conversion. After a preliminary characterization of antibody conjugated nanoparticle, I applied the FNPB-antibody to label the bacterial membrane. Bioimaging of the FNPB-antibody targeting bacterial membrane was measured by fluorescence microscopy. However, these are very preliminary results, more investigations are needed such as more characterizations of FNPB-antibody, and more experiments to confirm the microscopy experiments. The FNPB-antibody method is easier to be generated compared with the "sandwich" method. If the reproducibility can be proved, we can change the general anti-E.coli antibody to specific antibody for particular protein on bacterial membrane such as the antibody for protein controlling the growth of flagella. Using "nanoparticle-antibody" system to label this specific protein may allow us tracking the growth of flagella.

Conclusion

In this chapter, I presented two methods to target bacterial membrane with fluorescent nanoparticles: the "sandwich system" and the "nanoparticle-antibody" system.

In order to target E.coli's outer membrane with FNP -, a "sandwich system" was created (Figure 3.16). E.coli can be targeted with an antibody conjugated with biotin. The binding of biotin to streptavidin is one of the strongest non-covalent interactions known in nature.

After conjugating the biotin to nanoparticles and then with a streptavidin, we can use the particle based "sandwich system" to target the bacteria E.coli's outer membrane. In the "sandwich system", it takes three agents to link the FNP -to antibody which can target on bacteria (FNP --biotin-streptavidin-biotin-antibody). Even if it is proved by fluorescence (Figure 3.17A) and SEM (Figure 3.17B) that the "sandwich system" was successfully obtained, after applying it to label bacterial membrane, the targeting efficiency on bacteria is not very high. This may come from the several steps to form the sandwich. It was very difficult to obtain many cells targeted by particle. Moreover it was difficult to tell if the particle were really linked on the surface of bacteria.

In order to simplify this approach, we attempted to directly graft the anti-E.coli antibody to the FNPB but the characterization of the reaction is still underway. I applied the FNB-antibody to label the bacterial membrane, after a preliminary test, the particles conjugated with antibody can label bacterial outer membrane. However, this approach needs to be investigated more.

In the next chapter, I will describe several smaller fluorescent nanomaterials:

Fluorescent polymer chains (FPCs). I will show the interaction of FPCs with E.coli bacteria and the bioimaging applications of these FPCs.

Chapter 4 Fluorescent polymer chains (FPCs) for bioimaging of E.coli bacteria

In previous chapters, fluorescent nanoparticles were used for internal and surface bioimaging of E.coli bacteria. In chapter 1, we found that nanoparticles (65nm) are quite big and they can only be internalized in bacteria by electroporation. But it killed quite a lot of bacteria by this (40% survive). So, we wondered if it was possible to synthesize some small fluorescent nanomaterials which can enter easier inside bacteria. So, we had the idea to synthesis some polymer chains with a theoretical size of 4-6 nm.

In this chapter, I will present four different fluorescent polymer chains: negative green fluorescent polymer chain (GFPC -), neutral green fluorescent polymer chain (GFPC 0 ), negative red fluorescent polymer chain (RFPC -), positive red fluorescent polymer chain (RFPC -). I will discuss their physical characterizations and their interaction with E.coli bacteria for potential internal and membrane fluorescent labelling application.

Synthesis of fluorescent polymer chains (FPCs)

The synthesis of fluorescent polymer chains was performed in a novel one-pot process based on a reversible addition-fragmentation chain-transfer (RAFT) polymerization method:

a small RAFT agent and three different monomers: APEG, AA and BODIPY methacrylate-1 (BDPMA1) were dissolved in 1,4-dioxane at room temperature. After it was purged with nitrogen, the reaction of the mixture was initiated by simple heating to 80°C for 90 minutes.

After RAFT polymerization, the reaction was stopped by immersion of the flask in iced water.

Finally, after complete polymerization, a statistic and random copolymer (negative green polymer chain GFPC -) was formed: P(AA 9 -co-APEG 9 -co-BDPMA1 2 ) 20 -TTC-C 12 (Scheme 4.1) .

This polymer chain is negatively charged in water because the carboxylic acid group (-COOH, pKa around 4) will dissociate into H + cations and (-COO -) anions in water (pH 6.8-7.0). All four different FPCs have been synthesized (see experimental detail in Chapter 7, §7.2.4) with a similar architecture but different monomers in order to obtain products with a different charge and color: In GFPC°, the AA monomer was not added during the one-pot polymerization process in order to obtain a neutrally charged green polymer chain: P(APEG 17 -co-BDPMA1 3 ) 20 -TTC-C 12 (Scheme 4.2A). In order to make RFPC -, the BODIPY methylnaphthalene-2 (BDPMA2) monomer was used in order to obtain a red shift in

Characterizations of Green fluorescent polymer chains (GFPCs)

In order to characterize the green fluorescent polymer chains (GFPCs), samples were withdrawn regularly during polymerization, dried and analyzed by size exclusion chromatography (SEC, cooperation with Dr. Jutta Rieger ‡ ) in THF solution after methylation of the carboxylates. The SEC traces of copolymers obtained at different stages of the polymerization of GFPC -and GFPC° are displayed in which is slower compared with GFPC° (80%, Figure 4.1D). For GFPC -, a reaction plateau period was observed after 4 hours and after 8 hours it reached a final conversion of 70% for BDPMA1. While for GFPC°, the final conversion of BDPMA1 is around 95%. This proves that along the chain of GFPC°, there is about 25% more BDPMA1 converted which resulted in there is one fluorophore more than the one of GFPC -. The average molar mass, M n , increase linearly with monomer conversion and are close to the theoretical values. Polydispersity indexes, M w /M n , are low (<1.3, Table 4.1). The miniemulsion copolymerization of fluorescent monomer thus exhibits the features of a living well-controlled polymerization. M nth / g/mol [b] M nSEC / g/mol [c] M w /M The critical micelle concentration (CMC) of GFPCs in water was determined at 20°C or in LB medium at 37°C by dynamic light scattering (DLS, cooperation with Dr. Jutta Rieger § ) in order to determine the influence of temperature and growth medium. Different concentrations of GFPC -and GFPC 0 solutions were prepared and, the scattered light intensity at 90° was collected for these different samples. By plotting the intensity of scattered light as a function of GFPC -or GFPC 0 concentration, the CMC was determined at the intersection of the straight lines (Figure 4.

2).

The critical micelle concentration (CMC) of GFPC-is about 0.3mg/mL in water at 20°C and 0.4mg/ml in LB medium at 37°C. For GFPC 0 , the CMC is about 0.38mg/mL in water at 20°C and 0.5mg/ml in LB medium at 37°C (Table 4.2). The CMC of both GFPC-and GFPC 0 at 20°C is slightly higher than the one at 37°C, this is because the increase in temperature will § LCP laboratory (UMR 7610), UPMC Univ Paris 06, France cause entropy increases which induce the structure to break apart, and thus a higher concentration of surfactants is needed to form micelles. The CMC of GFPC -is always lower than the one of GFPC 0 , this is perhaps because GFPC -is charged and the polar head of GFPC - are better located and organized than GFPC 0 which is more unorganized, randomly distributed in solution. The concentration of GFPCs that will be used in experiments with E.coli bacteria should be below the CMC so that the GFPCs are not aggregating to form micelles when incubated with E.coli bacteria. 4.3. The BDPMA1 monomer in toluene shows the expected spectra for BODIPY fluorophores [START_REF] Loudet | BODIPY dyes and their derivatives: syntheses and spectroscopic properties[END_REF] with an intense band in the visible region located at 528 nm (corresponding to transition) and a vibrational shoulder at higher energy. A second, less intense band is located in the UV region at around 380 nm. The maximum of fluorescence emission is found at 540 nm, exhibiting a Stokes shift of 421cm -1 (12nm).

The GFPCs exhibit very similar absorption and fluorescence spectra with a maximum absorption at 528 nm and a maximum of fluorescence emission at 548 nm. Both GFPC -and GFPC° are identical in shape and position, this shows that the addition of the AA monomer doesn't change the spectrum of GFPCs. The Stokes shift of the nanoparticles (691cm -1 , 20 nm) is slightly higher than the monomer. The red shift of the fluorescence emission is observed from the free monomer to the monomer copolymerized in a rigid environment 180 .

One can also notice that the GFPCs absorption spectra exhibit a more intense vibrational shoulder which might also result from the BODIPY confined in a rigid polymer matrix (Figure In water the fluorescence quantum yield of GFPC 0 is 29% while the one for GFPC -is 20%. The fluorescence quantum yield of BDPMA1 in toluene is 70%. This difference may be due to the aggregation of BODIPY along the same polymer chain. To confirm this hypothesis, I analyzed the GFPC 0 and GFPC -spectra in toluene (Figure 4.5). The fluorescence quantum yield is 78% for GFPC 0 and 74% for GFPC -(Table 4.3). Since toluene is a good solvent for this kind of polymer chains, they will be reformulated in it. The spectra of GFPCs in toluene are identical with the one of BDPMA1. So, there will not be any aggregation of BODIPY along the same polymer chain. That is probably why the quantum yield of GFPCs in toluene is higher than the GFPCs in water.

Toxicity assessments of GFPCs on E.coli

Having synthesized and characterized the GFPCs, I next assessed the toxic effect of GFPCs on E.coli. In order to do this, the growth rates of bacteria in the presence of GFPCs were compared with the growth rates of E.coli bacteria alone. The growth rate was measured by using a 96-well plate reader (Perkin Elmer Victor3 1420 Multilabel Plate Counter). The GFPCs concentration used was 6.6x10 -5 M. The concentration of GFPC 0 or GFPC -(0.015-0.025 mg/mL) in all samples is below the CMC (0.3-0.5 mg/mL) of GFPCs, means that GFPCs are not aggregating to form micelles when incubated with E.coli bacteria.

Here, E.coli bacteria with GFPC° or GFPC -were incubated in the 96-well plate in plate reader at 37°C in the dark overnight. A control with only bacteria was also prepared. Every nine minutes the growth rate of the cells was monitored by measuring the optical density at 600nm (Figure 4.6). For over ten hours incubation, the growth curves of bacteria with GFPCs were practically the same as the growth curve of bacteria alone both in LB medium and M9 medium. This means that this concentration of GFPCs has no toxicity effect on bacterial growth in a long time scale independently of the charge of the GFPCs. After make sure that

GFPCs have no toxic effect on E.coli bacteria, and then I investigated their interaction with bacteria by imaging.

A B

GFPCs for internal bioimaging application of E.coli bacteria 4.4.1 Bioimaging of E.coli interacting with GFPCs

Characterization by flow cytometry

After having shown that both GFPC -and GFPC 0 have no toxicity effect on E.coli bacterial growths, the GFPCs were tested for bioimaging application with E.coli. As the theoretical size of the GFPCs is around 5 nm, they should enter inside bacteria just by incubation 183 .

In order to measure whether the GFPCs can enter inside E.coli successfully, the LB growth medium with an initial E.coli bacterial concentration of 5x10 7 CFU/mL (OD=0.5) was placed inside an Eppendorf Safe-Lock Tube in the presence of GFPCs. The samples were incubated at 37°C in the dark shaking for one hour and washed for twice with PBS to remove free GFPCs. The cells were then fixed and then washed three times with PBS and resuspended in PBS before flow cytometry measurements and bioimaging experiments.

Accurate measurements of live, dead, total bacteria and fluorescent cells percentage (Figure 4.7) after incubation of E.coli with different concentration of GFPCs (6.6x10 -4 M, 6.6x10 -5 M, 6.6x10 -6 M) were obtained by Flow Cytometry.

After incubation of E.coli with GFPC 0 , the percentage of total fluorescent cells increased when increasing the concentration of GFPC 0 , and it reached the plateau when the concentration is 6.6x10 -5 M (Figure 4.7A). For the fluorescent cells which are alive, the percentage increases at the beginning, and reaches the plateau at the concentration of 6.6x10 -5 M as well. The percentage of fluorescent live cells could reach at about 80%. From the plate reader experiments described in last section, GFPCs have no toxic effect on E.coli bacteria, so the 20% loss of bacteria may come from sample preparation (washing, fixing process…).

After incubation of E.coli with GFPC -, the percentage of total fluorescent cells increases quite fast from the beginning and it remains at the plateau no matter what the concentration is (Figure 4.7B). For the live fluorescent cells, the percentage remains at about 80% from the beginning. As well as the GFPC 0 , GFPC -also has almost no toxicity effect on E.coli's growth.

Comparing GFPC 0 and GFPC -interacting with the E.coli bacteria (Figure 4.7), we found that GFPC -will associated to the E.coli at a lower concentration. However GFPC 0 can also reach a high percentage of cell labeling if we increase its concentration. This is a surprising result because GFPC -is negatively charged, and GFPC 0 is neutral, and the membrane of E.coli bacteria is also negatively charged, 199 thus one could expect that GFPC -should have lower probability to associate with cells, which is not the case from the experiments results. Taking into account the percentage of live cells and live fluorescent cells, 6.6x10 -5 M proved to be a suitable concentration for the following experiments.

Additionally, In

Bioimaging of E.coli interacting with GFPCs

E.coli's interaction with GFPC 0 and GFPC -was measured by epifluorescence microscopy.

The fluorescence images of E.coli alone (Figure 4.8A) show the auto-fluorescence of E.coli.

After incubation of E.coli with GFPCs, under the microscope the E.coli cells becomes green, it seems that the GFPCs entered into the cells, ( previous section , we found out that the quantum yield of GFPC 0 (29%) is higher than GFPC - (20%) in water and brightness of GFPC 0 is twice as high compared with GFPC -(Table 4.3), from image, it seems that under the same condition more GFPC -will be incorporated into E.coli than GFPC 0 . However, until now, it is difficult to tell if these GFPCs enter inside bacteria or attach on the membrane of the bacteria. From the flow cytometry characterization, I determined that 80% of the cells are alive and fluorescent after incubation with the GFPCs, and from the fluorescence microscopy images, we can also conclude that GFPCs (4-6 nm) can associated with the bacteria, especially for the negatively charged GFPC -.For the GFPC 0 , each bacterium "takes" a different amount of polymer chains, resulting in bacteria of different brightness. However for the GFPC -, the fluorescence of the bacteria is more homogeneous. Since E.coli bacteria incorporated more easily the GFPC -resulting in brighter and better quality fluorescent images after incubation, in the following experiments, I decided to concentrate on GFPC -.

A B C

Emission quenching of GFPC -with Methylene Blue (MB)

Characterization of GFPC -quenching with Methylene Blue (MB)

In order to make sure that the GFPCs were really inside E.coli after incubation not on the membrane; methylene blue was used during the bioimaging experiments. Methylene blue is a biocompatible dye that can quench GFPCs, the idea is similar as using MB to quench FNP -which described in Chapter 2 ( §2.4.3).

Here we take GFPC -as an example since GFPC -has a better ability to label bacterial cytoplasm from previous experiments. Titration experiments were prepared to characterize the process of GFPC -quenching with MB. A GFPC -solution was placed inside a cuvette and methylene blue solution was added continually inside the GFPC -solution and emission spectrum was taken each time until the emission of GFPC -stopped decreasing.

The fluorescence and absorption spectroscopy analysis of GFPC -and methylene blue are displayed on In order to understand the mechanism of the quenching process, the data was plotted according to Stern-Volmer equation in 7.4 in Chapter 7). However, the plot is still not linearly.

However, the plot is still not linearly. The modified Stern-Volmer plot is only suitable for conditions that contain two populations of fluorophores (one accessible and one inaccessible). In the case of GFPC -, the system seems more complex. We may postulate the existence of some dimers or trimers or n-mers aggregates which form a complicated "solution". So far we can't explain the dynamics by modified Stern-Volmer plot.

A B

Bioimaging after quenching with Methylene Blue (MB)

After characterization of the quenching process of GFPC -, methylene blue was used in the bioimaging experiments of E.coli interacting with GFPC -. E.coli's interaction with GFPC - after the addition of methylene blue was also measured by epifluorescence microscopy.

After adding methylene blue, the GFPCs outside the cells were quenched by the methylene blue, thus on DRAQ5 is a red fluorescent cell-permeable DNA dye that is biocompatible. [START_REF] Smith | DRAQ5 labeling of nuclear DNA in live and fixed cells[END_REF] The chemical structure and spectral characteristics of DRAQ5 are shown in PKH26 was first tested on E.coli for the ability to label bacterial membrane, As show in It is important to notice that only a few bacterial membranes were successfully labelled by PKH26 while almost all the bacterial cytoplasm were labelled successfully by GFPC -. I overlapped the two images together (Figure 4.18D), and it appeared that the yellow part and the green part are not influenced by each other, and from the image it is clear that the bacterial cytoplasm is green while the membrane is yellow. This is additional evidence to show that GFPC -can label the bacterial cytoplasm. At the same time these results indicate that GFPC -are quite compatible with different types of commercial dyes that could be used to multi-color imaging.

Characterizations of Red fluorescent polymer chains (RFPCs)

The previous results showed that, GFPC -could be used to label the bacterial cytoplasm but not the DNA complex which is negatively charged. It would be interesting to synthesize some positively charged polymer chains that could label bacterial DNA for bioimaging. However, the maximum band of fluorescence emission at 601nm (RFPC + ), 588nm (RFPC -)

indicates a red shift of RFPC + compared with RFPC -. Moreover the emission band of RFPC + is broader than the one of RFPC -. These two facts brought together may indicate a higher aggregation state of RFPC + in water. It is observed that the RFPCs absorption spectra exhibit a more intense band located in the UV region which may come from the light scattering by the polymer chains aggregations. This is more pronounced for RFPC + than for RFPC -. This is also in agreement with a higher aggregation state in RFPC + (Figure 4 The fluorescence quantum yield of BDPMA2 in ethanol is 53% which shows about a 20% decrease from the one of BDPMA1 (70%). The additional aromatic moieties on the pyrrole of the BODIPY monomer not only shift fluorescence emission to red but also make the monomers being congested [START_REF] Grazon | Élaboration de nanoparticules fluorescentes à base de BODIPY par polymérisation RAFT en miniémulsion : synthèse, caractérisation et fonctionnalisation de surface[END_REF] . In water the fluorescence quantum yield of RFPC + is 12% while the one for RFPC -is 10%. The decrease may be due to the aggregation of BODIPY along the same polymer chain. To confirm this, I analyzed the RFPC + and RFPC -spectra in ethanol (Figure 4.22). The quantum yield is 32% for RFPC + and 31% for RFPC -(Table 4.4). The spectra of RFPCs in ethanol still show intense band in UV region indicating that there were still aggregations inside the solution. That is probably why the quantum yield of RFPCs in ethanol is higher than the RFPCs in water but still lower than BDPMA2 in ethanol.

Toxicity assessments of RFPCs on E.coli

Having synthesized and having obtained a preliminary characterization of the RFPCs, I next assessed the toxic effect of RFPCs on E.coli. In order to do this, the growth rates of bacteria in the presence of RFPCs were compared with the growth rate of E.coli bacteria alone. The growth rate was measured by the plate reader (Perkin Elmer Victor3 1420

Multilabel Plate Counter). The RFPCs concentrations used was 2x10 -10 M.

Here, E.coli bacteria with RFPC + or RFPC -were incubated in the plate reader at 37°C in the dark overnight. One control with only bacteria was also prepared. Every nine minutes the growth rate of the cells was monitored by measuring the optical density at 600nm by the plate reader (

RFPCs for external bioimaging application of E.coli bacteria 4.7.1 Bioimaging of RFPCs labelling the membrane of E.coli

After having shown that both RFPC + and RFPC -have no toxic effect on E.coli bacterial growth, RFPCs were tested for bioimaging application with E.coli. Taking into consideration the results from GFPCs, we expected that RFPC + would enter inside bacteria and label the DNA which is negatively charged, and RFPC -would enter inside bacteria and label the cytoplasm like the GFPC -did. However, the results are quite different from those expected.

E.coli's interaction with RFPC + and RFPC -was measured by epifluorescence microscope (TRITC filter). The auto-fluorescence of E.coli alone is shown in Figure 4.24 which indicates that there is no red auto-fluorescence of E.coli.

Multi-color Imaging application of GFPC -and RFPC +

Since RFPCs can label the bacterial membrane, I next tested the multi-color internal and external bioimaging of E.coli bacteria with GFPC -and RFPC + together. RFPC + was added to label the bacterial membrane after GFPC -was introduced inside bacteria in order to prevent the possibilities that RFPC + may block the GFPC -to enter.

Comparison of PKH26 and RFPCs

From the previous experiments we can conclude that our novel highly water-soluble red fluorescent polymer chains RFPCs appear to label bacterial membrane very efficiently.

Compared with the commercial membrane dye, PKH26, used in previous experiments, RFPCs can label bacterial membrane in a clear and homogeneous way, while for PKH26, there were some holes on the membrane labelling ( The staining concentration for RFPCs during the previous experiments described above is 2x10 -10 M for 1x10 8 cells/mL. While the label concentration for PKH26 is of 2x10 -6 M for 1x10 8 cells/ml which is 10000 times higher compared with the polymer chains. This means that we can easily use 10000 times less polymer chains to label the membrane of bacteria From the experiments described above, a decrease in the amount of PKH26 resulted in a decrease in staining efficiency. In

Conclusion

In this chapter, I presented four different fluorescent polymer chains (FPCs): neutral and negatively charged green fluorescent chains (GFPC°, GFCP -), positively and negatively charged red fluorescent chains (RFPC + , RFPC -) and their bioimaging applications on E.coli bacteria (Figure 4.31).

GFPCs have been shown to be nontoxic on bacterial growth in a long time scale, independently of the charge and of the growth medium. Since E.coli bacteria incorporated more easily the GFPC -resulting in brighter and better quality fluorescent images, I therefore decided to concentrate on GFPC -.

I also developed the multi-color application of GFPC -. DRAQ5 is introduced to label bacterial DNA while at the same time the cytoplasm of bacteria is labelled by GFPC -. It is found that GFPC -could only label the bacterial cytoplasm but not the DNA. Another dye PKH26 is also introduced to label bacterial membrane while GFPC -is used for labelling the cytoplasm.

Chapter 5 Fluorescent pH-sensitive nanoparticles (FANPs) for rapid detection of E.coli growth

Rapid detection of bacterial growth is an important issue in the food industry and for medical research. In this chapter I will present a novel kind of fluorescent pH-sensitive nanoparticles (FANPs) that can be used for the rapid and accurate real-time detection of Escherichia coli growth. These organic particles are designed to be non-toxic and highly water-soluble. Here we show that the coupling of pH sensitive fluoresceinamine to the nanoparticles results in an increased sensitivity to changes in pH within a physiologically relevant range that can be used to monitor the presence of live bacteria. In addition, these FANPs do not influence bacterial growth and are stable over several hours in a complex medium and in the presence of bacteria. The use of these FANPs allows for continuous monitoring of bacterial growth via real-time detection over long time scales in small volumes and can thus be used for the screening of a large number of samples for high-throughput applications such as screening for the presence of antibiotic resistant strains. This part of work has been submitted.

Synthesis of fluorescent pH-sensitive nanoparticles (FANPs)

Synthesis of polystyrene nanoparticles (PSNPs)

The pH-sensitive nanoparticles presented here have been elaborated from polystyrene nanoparticles (PSNPs) as recently reported 180 . The one-pot synthesis is based on a reversible addition-fragmentation chain-transfer (RAFT) polymerization in miniemulsion conditions using a surfactant-free one-pot phase inversion process (Scheme 5.1). Briefly, the particles were obtained from a hydrophilic chain PEO-b-PAA (poly(ethylene oxide)-blockpoly(acrylic acid)) macroRAFT agent. These hydrophilic chains were first extended with a hydrophobic block by a bulk polymerization of styrene (S). The polymerization was stopped before total consummation of styrene and, by adding basic water, droplets of the remaining styrene covered by the amphiphilic copolymer PEO-b-PAA-b-PS were obtained. A homogeneous miniemulsion was formed by ultrasound treatment. Finally, the RAFT polymerization was reinitiated by simple degasing with argon and heating to 80°C. After repolymerization, a core-shell structure negatively charged nanoparticle was obtained with polystyrene in the core and poly(ethylene oxide)-block-poly(acrylic acid) in the shell. The obtained PSNP1 was well-defined in size with a mean hydrodynamic diameter of 70 nm 27 (Table 5.1).

Scheme 5.1 Synthetic scheme employed for the synthesis of the PSNP1. In order to obtain a stable colloidal dispersions, a poly(acrylic acid) block is used in the particle design in order to have both electrostatic and steric repulsion. In addition, the introduction of carboxylic acid groups in the particle's shell is helpful for post-functionalizing of the PSNPs and the preparation of pH-sensors. Furthermore, in order to test the effect of the architecture of the hydrophilic chains on the pH-dependence of fluorescence, a PSNP2 with a different shell architecture compared with PSNP1 was designed. In this case, the PEOb-PAA shell was replaced by a randomly distributed copolymer of acrylic acid (AA) and poly (ethylene oxide) methyl ether acrylate (PEOA) as P(PEOA-co-AA). It was found that PSNP2 nanoparticles have a similar hydrodynamic diameter compared with PSNP1 (Table 5.1).

Grafting pH sensitive fluorophores on the PSNPs

Fluoresceininamine was selected as a pH-sensitive molecule because it has an intense absorption band in the visible. Even though it can adopt four different forms (dianion, anion, neutral and cation) depending on the pH, it is only highly fluorescent in its dianionic form 201 . Furthermore, the pKa of the anion/dianion couple is about 6.4, which is suitable to measure changes in pH in biological media. The grafting of amino-functionalized molecules on carboxylic groups is a standard procedure 202 . The functionalization of PSNPs with fluoresceinamine (FA) was performed at pH=8 and 4 °C. The grafting of FA on PSNP1 and PSNP2 was carried out using 0.5 and 1 equivalent of FA per acrylic acid unit, respectively (Scheme 5.2). In all cases, the grafting did not change the hydrodynamic diameter of the nanoparticles significantly. The hydrodynamic of the fluoresceinamine conjugated nanoparticles (FANPs) remains around 50-70 nm (Table 5.2). Scheme 5.2 Preparation of pH nanosensors (FANP1 and FANP2) from PSNP1 and PSNP2.

Characterization of FANPs

Spectroscopic properties of the FANPs

Absorption and fluorescence spectroscopy analysis of FANP1 and FANP2 are shown in Where I is the fluorescence intensity of the sample and I A and I B are the fluorescence intensities in the acidic and basic forms respectively [START_REF] Valeur | Molecular Fluorescence[END_REF] . The pKa for both types of nanoparticles is around 6.5 and a linear dependence of the fluorescence intensity was observed between pH 5.5 and 7.5. Both the pKa and the linear range are in agreement with the one of the free FA dye 205 , therefore it seems that the grafting does not change the pH dependent fluorescent properties of FA. This is the useful pH range for the study of most biological systems. Compared to approaches reported previously 176,206 , in this case the linearity of the signal covers a wider and more suitable range of pH values for detecting bacteria. The amplitude of the fluorescence intensity change between these two pH values is around 7-fold which is 3 times higher than previously reported 176,207 . These properties will thus allow a sensitive and precise determination of the change in pH in biological systems. pH values: 7.37, 7.07, 6.94, 6.41, 5.84, 4.95, 4.41, 3.85, 3.01. D: Change in fluorescence intensity of FANP2 at 520nm as a function of pH. The data was fit using the Henderson-Hasselbach equation with pKa=6. 53. pH values: 7.80, 7.14, 7.01, 6.55, 6.27, 5.85, 5.07, 4.57, 4.00.

Photostability and reversibility assessments of FANPs

It is widely known that the main drawback of fluorescent sensors is the bleaching of the fluorophore over long time periods. The photostability of the FANPs was measured to determine whether photo-bleaching occurs during extended measurement of fluorescence during the incubation in the growth medium. FANPs were added to the modified M9 minimal medium at room temperature or 37°C. Every 30 minutes a fluorescence emission spectrum was recorded (Figure 5.3). It was found that over four hours there is no fluorescence decrease for both FANP1 and FANP2.

Toxicity assessments of FANPs on E.coli

In order to assess the toxic effect of FANPs on E.coli, the growth rate of bacteria in the presence of FANPs was compared with the growth rate of E.coli bacteria alone. The growth rate was measured by absorbance measurement (OD@600nm) in a 96-well plate reader. The FANPs concentrations used were 8.2x10 -7 M, 4.1x10 -7 M and 8.2x10 -8 M.

E.coli bacteria with different concentrations FANP1 or FANP2 were incubated in the plate reader at 37°C in the dark overnight. A control with only bacteria was also prepared.

Detection of E.coli growth with FANPs

Real-time detection of E.coli growth

Growth of E.coli bacteria with glucose results in the production of CO 2 and acetic acid and, therefore, a decrease in pH of the growth medium. Having synthetized and characterized the pH-sensitive nanoparticles, we next tested whether they could be used to precisely monitor the pH change due to bacterial growth. In order to measure the change in fluorescence intensity of FANPs in the presence of E.coli bacteria, the growth medium with an initial pH of 7.0 and an initial E.coli bacterial concentration of 2x10 7 CFU/mL(OD=0.2) was placed inside a cuvette in the presence of FANPs. E.coli bacteria samples with the FANPs were incubated at 37°C in the dark. Every 30 minutes the growth of the cells was monitored by measuring the optical density at 600 nm using a spectrophotometer (Figure 5.7E, F).

When the bacteria began to grow, the pH of the growth medium started to decrease. The pH of the samples was monitored at several time intervals with a pH meter. When the bacterial concentration increased to 1.1x10 8 CFU/mL (OD=1.1), the pH of the growth medium decreased to 5.8. At the same time, the emission spectra of the samples were measured. As

Sensitivity assessments

A successful pH nano-sensor for the detection of E.coli growth should be both rapid and sensitive to small changes in bacterial concentrations. We therefore performed assays in order to assess the sensitivity of FANPs compared with the one of the same concentration of free fluoresceinamine (FA) molecules using a lower starting concentration of bacteria. FANP1, FANP2 or FA molecules were added to the growth medium with an initial concentration of E.coli bacteria of 3-6x10 5 CFU/mL (OD=0.003-0.006) inside a cuvette and were incubated at As mentioned above, the growth of the cells was monitored using a spectrophotometer that reads the optical density (OD@600nm) of the samples every 30 minutes (Figure 5.8). From the time at which the bacteria started to grow until two hours of growth, it was found that this small amount of bacteria couldn't be measured accurately by a measurement of the change in optical density. In addition, there were large fluctuations in the signal in the first hour of growth, and as the bacterial number increased, the measurements of spectrophotometers became more reproducible. However, as shown in These results show that these pH-sensitive nanoparticles are not only more sensitive than a molecular sensor such as the FA molecules but also more sensitive than typical OD measurements.

High-throughput screening applications

Screening of cell growth in a 96 well plate

Since the FANPs can be used to measure the change in pH resulting from the metabolic activity of live bacteria, they can be used to discriminate whether the bacteria have died or have a decreased growth rate due to the fluorescence intensity change caused by metabolism. This approach can thus be more sensitive compared with typical OD meters and can be performed in complex media. We next tested the potential ability of FANPs for highthroughput screening of E.coli bacteria.

In order to measure the fluorescence evolution of FANPs when incubated with smaller volumes of bacteria, E.coli were incubated in a 96-well plate with different concentrations (8.2x10 -7 M, 4.1x10 -7 M and 8.2x10 -8 M) of FANP1 or FANP2 and placed in a plate reader at 37°C in the dark overnight. A control with bacteria without nanoparticles was also prepared.

The fluorescence intensity of each well was monitored every 6 minutes by using a F485/14

filter for excitation and a F535/40 filter for emission (Figure As a comparison the amount of FANP1 used in the cuvette experiment was the highest of the three used here, 8.2x10 -7 M.This shows that even after a ten-fold decrease in the concentration of FANP1 (8.2x10 -7 M8.2x10 -8 M), one can still detect bacterial growth rapidly and accurately. Because of their stability and sensitivity these FANPs could be used as a novel indicator for screening of potential growth inhibition experiments.

High-throughput screening of bacterial growth inhibition by antibiotics

After demonstrating that FANPs can be used for measurement of E.coli bacteria growth in small volumes, we wanted to determine whether this approach could be used for Moreover, from the growth curve, one can notice that after about ten hours of incubation bacterial growth, as measured by the OD, reached the stationary phase, indicating that cell growth slowed down and that bacteria grew and died at the same rate, resulting in a constant number of bacteria. However, by measuring the OD it is not possible to measure the presence of bacterial growth once the population has reached stationary phase. On the other hand, the increase in the change in fluorescence at these later time points allows for a measurement of the amount of ongoing metabolism of E.coli. The same results were obtained for FANP2 (Figure 5.13). As anticipated, reducing the dose of antibiotics reduces its inhibitory effect, and the changes in cell metabolism can be measured reliably and accurately by using FANPs. In order to test whether these FANPs could be used to monitor bacterial growth on a solid support, they were embedded in agar with growth medium, as used traditionally to follow the formation of bacterial colonies. This result indicates that these particles could be used to screen for bacteria that grow selectively on solid media.

Imaging of E.coli growth

Conclusion

The novel pH-sensitive nanoparticles described herein have distinct features that make them suitable for the development of biosensors including brightness, stability in growth medium, non-toxicity, high water solubility, high sensitivity, and ease of bioconjugation and incorporation into a biocompatible matrix. They can rapidly and accurately detect bacterial growth by detecting the change of pH resulting from cellular metabolism. Due to the brightness of these nanoparticles, they could be used to develop new systems for quantitative biosensor applications for microfluidic devices. One of the important advantages of this approach is that a change in pH can be easily monitored either by a simple change in fluorescence at a specific wavelength or even by eye for a qualitative result in either a liquid or solid support. These FNAPs can be easily synthesized with low cost and can be used in any laboratory without additional expensive instruments or operators.

Moreover, these particles allow for continuous monitoring for long time scale experiments and can be used to monitor a large number of samples for high-throughput screening applications. For example they can be used to measure the metabolic activities of bacteria as an indicator of their susceptibility to antibacterial drugs, as shown here, or for the screening and discovery of novel growth inhibitors.

Chapter 6 General conclusion and perspectives

The main aim of my PhD research project was to study fluorescent nanomaterials for bioimaging and biosensing on E.coli bacteria.

Nine different fluorescent nanomaterials were synthesized, characterized and used as fluorescent agents for biological experiments with E.coli bacteria: Fluorescent nanoparticles (FNP -, FNP + , FNP-PEG + ), Fluorescent polymer chains (GFPC -, GFPC 0 , RFPC -, RFPC + ) and fluorescent pH-sensitive nanoparticles (FANP1 and FNAP2). All these fluorescent nanomaterials have been proved to be nontoxic to E.coli bacteria.

The negatively fluorescent nanoparticles FNP -have a core-shell structure with around 3000 BODIPY fluorophores copolymerized in the core and hydrophilic polymer chains in the shell, the size of the particle is around 65nm. Thanks to the thousands of fluorophores inside the core, FNP -brightness was very high (10 7 M -1 .cm -1 ). It is approximately 1000 times brighter than traditional organic dyes such as BODIPY and Fluorescein. They are also very stable due to the fluorophores protection in the hydrophobic core. By coupling reactions with amine, two different positively charged fluorescent nanoparticles are obtained: FNP + and FNP-PEG + . These fluorescent nanomaterials described in the manuscript have proved to be ideal tools for biolabeling and bioimaging applications in life science due to their high brightness, high photostability and rich functionalization ability.

In order to study the internal bioimaging of E.coli with FNPs, electroporation was performed to insert FNPs inside bacteria (Figure 6.1A). Bioimaging experiments indicated that E.coli bacteria interacted with fluorescent nanoparticles. Methylene blue was introduced to quench the fluorescence of particles outside the bacteria, the fluorescence of particles inside bacteria remained thus proving that particles are successfully inserted inside bacteria. However, very few bacteria survived after electroporation. This is a strong drawback for internal bioimaging application. New strategies were needed to apply the particles to bioimaging of E.coli. One was to use the particles for external bioimaging of E.coli.

On the other hand, if these FNPs bind selectively to dead bacterial cells they could be used to measure the toxicity of potential antibiotic molecules using a small number of cells for drug screening application.

Thanks to the poly acrylic acid groups all around the particle, biological molecules (e.g. biotin, antibody…) could be conjugated on the surface of the particles (Figure 6.1B). In order to study the external bioimaging of E.coli with FNPs, a "sandwich system" was created to target the bacterial membrane. E.coli was targeted with an antibody conjugated with biotin.

After conjugating the biotin to nanoparticles and then with a streptavidin, I used this particle based "sandwich system" (FNP --biotin-streptavidin-biotin-antibody-bacteria) to target the E.coli's outer membrane. In the "sandwich system", three agents are needed to link the FNP - to the antibody which can target the bacteria. Even if it was shown by fluorescence and SEM images that the "sandwich system" was successfully obtained, the targeting efficiency on bacteria was not high enough for further application. In order to simplify this approach, I attempted to directly graft the anti-E.coli antibody to the surface of fluorescent nanoparticles but the characterization of the reaction is still underway. I applied the "nanoparticle-antibody" system to label the bacterial membrane, after a preliminary test, the particles conjugated with antibody could label bacterial outer membrane. Fluorescent nanoparticles with a size of 65nm proved to be too big and they could only be internalized in bacteria by electroporation. Smaller fluorescent nanomaterial: fluorescent polymer chains (4-6nm) were thus synthesized and investigated for bioimaging on E.coli bacteria.

Green fluorescent polymer chains (GFPCs) could easily enter inside bacteria by simple incubation (Figure 6.1C). Particularly, flow cytometry experiments showed that GFPC -could label all the bacteria. I also developed a multi-color application of GFPC -. DRAQ5 was introduced to label bacterial DNA while at the same time the cytoplasm of bacteria was labelled by GFPC -. It was found that GFPC -could only label the bacterial cytoplasm but not the DNA. Another dye PKH26 was also introduced to label bacterial membrane while GFPC - was used for labelling the cytoplasm. One of the important advantages of this approach is that a change in pH can be easily monitored either by a simple change in fluorescence or even by eye for a qualitative result in either a liquid or solid support. These FNAPs can be easily synthesized with low cost and can be used in any laboratory without additional expensive instruments or operators. Moreover, these particles allow for continuous monitoring for long time scale experiments and can be used to monitor a large number of samples for high-throughput screening applications. For example we used them to measure the metabolic activities of bacteria as an indicator of their susceptibility to antibacterial drugs.

Various potential approaches may need to be investigated in the future. Eukaryotic cells are much bigger compared with bacteria, FNPs may be internalized by cells through the process of endocytosis. Due to their high brightness and high photostability, FNPs can be useful tools for single-particle tracking experiments inside eukaryotic cells to understand dynamics in cells 209 . FNPs based "Sandwich system" and "Nanoparticle-antibody" system can be used to study specific membrane protein dynamics 210 on bacteria or eukaryotic cells in the future.

For example, by changing the general anti-E.coli antibody to the specific antibody for membrane protein which control the growth of flagella, nanoparticles can label particularly on this membrane protein and may allow tracking the growth dynamics of flagella 211 .

Besides bioimaging application, in the future, GFPC -can also be used to distinguish and quantify live and dead bacteria with a combination of propidium iodide (PI) by flow cytometry. GFPC -can label all bacteria while PI can label only dead cells, the viability is measured according to the proportion of bound strains by GFPC -and PI 200 . RFPCs have shown great potential for membrane labelling, in the future, they can be investigated for labeling of the membrane of eukaryotic cells, and study the cells' division by microscopy.

FANPs also possess great potential for various approaches, they can be used to develop new systems for quantitative biosensor applications for microfluidic devices or for the screening and discovery of novel growth inhibitors. These particles could also be used to study the change in metabolism in different biological systems such as during the formation of biofilms 212 , in the event of skin infections 213 or even to detect the presence of tumour cells during surgery due to their acidic environment 214 .

Chapter 7 Experimental methods and materials

Materials

Acrylic acid (99%, Sigma-Aldrich, AA), poly(ethylene oxide) methyl ether acrylate (Mn=480 g/mol, Sigma-Aldrich, APEG), 4,4'-azobis(4-cyanopentanoic acid) (Sigma-Aldrich, ACPA), 2-methyl-2-[(dodecylsulfanylthiocarbonyl)sulfanyl] propanoic acid (> 97%, Strem, TTCA), 2-(dimethylamino)ethyl acrylate (98%, Sigma-Aldrich, DMAEA), 1-ethyl-3-(3dimethylaminopropyl) carbodiimide hydrochloride (>98%, Fluka, EDC.HCl), ethylenediamine (>99%, Sigma-Aldrich), Poly(ethylene glycol) diamine (Sigma-Aldrich), Fluoresceinamine, isomer I (Sigma-Aldrich, FA), ethanolamine (>99%, Sigma-Aldrich, EtA), citric acid (99.8%, Carlo Erba), sodium phosphate dibasic dihydrate (>99.5%, Sigma-Aldrich), paraformaldehyde (Sigma-Aldrich, PFA), Poly(ethylene glycol) 2-aminoethyl ether biotin(average Mn 2300 (n~45), Sigma-Aldrich), Anti-E.coli antibody ab13627(IgG, abcam), Anti-E.coli antibody (Biotin) ab20640 (IgG, abcam), Anti-rabbit IgG (H+L), F(ab')2 Fragment (Alexa Fluor® 488 Conjugate) #4412 (Cell Signaling Technology), Streptavidin (Sigma-Aldrich), Formaldehyde (Sigma-Aldrich, 36.5-38%) were used as received. Chloramphenicol, kanamycin, and ampicillin antibiotics were obtained from Sigma-Aldrich and used as received.

2,2'-Azobis(2-methylpropionitrile) (AIBN, Sigma, 98%) was recrystallized from chloroform and few drops of petroleum ether. Styrene (Sigma Aldrich, ≥99%) and Dichloromethane (97%, Fluka, DCM) were distilled under reduced pressure. Distilled water was used for nanoparticles synthesis, the pH of water was adjusted with aqueous sodium hydroxide solution (1M) to a value comprised between 12 and 12.5. Solvents (Carlo Erba) were of synthetic grade and purified according to standard procedures. Silica gel 60A ˚ (70-200mm porosity) was bought from SDS. Deionized water (15M.cm at 20°C) was prepared with a Milli-Q system (Millipore). Polyethylene glycol, PAA: Polyacrylic Acid, TTC: Trithiocarbonate), copolymers as macroRAFT agent in a one-pot phase inversion process. In a septum-sealed flask, PEO-b-PAA macroRAFT (3.9x10 -5 mol, 137 mg) is dissolved with a mixture of styrene (6.45x10 -3 mol, 671 mg), AIBN (Azobisisobutyronitrile) (2.0 mg, 1.24x10 -5 mol) and BDPMA1 (6.68x10 -5 mol, 31 mg). The mixture is purged with argon for 30 min in an ice bath, and then placed in an oil bath thermostated at 80°C to initiate polymerization. After 70 min, the reaction is stopped by immersion of the flask in iced water (Scheme 7.1).

To the cold organic mixture, 5mL of basic water (pH=12.5) is added. An ultrasonic horn (Bandelinelectronics, SonoplusHD2200) is then placed in the biphasic mixture cooled down in an ice bath and powered at 130 W for 10 min. After the miniemulsion formation, the pH decreased to 11. The miniemulsion is purged with argon for 30 min in an ice bath, and then placed in an oil bath thermostated at 80°C to re-initiate polymerization.

Conversion of FNP -to FNP + and FNP-PEG + (Scheme 7.2)

Coupling Reaction 1 (FNP + ) Ethylenediamine (10.8 mg, 179.4 µmol) and fluorescent nanoparticle (FNP -) (2.58 mg, 35.9 µmol) are dissolved in distilled water (4 mL). A mixture of EDC.HCl {1-ethyl-3-(3dimethylaminopropyl) carbodiimide hydrochloride } (27 mg, 143.5 µmol) in water is added to the previous solution which was subsequently stirred at 4°C for 24h and then dialyzed against water (pH=5-6) to remove the excess ethylenediamine. The water (1L) surrounding the dialysis tube was changed once a day during the process for one week.

Coupling Reaction 2 (FNP-PEG + ) Poly(ethylene glycol) diamine (39.5 mg, 179.4 µmol) and fluorescent nanoparticle (FNP -) (2.58 mg, 35.9 µmol) were dissolved in distilled water (4 mL). A mixture of EDC.HCl {1ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride } (27 mg, 143.5 µmol) in water is added to the previous solution which was subsequently stirred at 4°C for 24h and then dialyzed against water (pH=5-6) to remove the excess of poly(ethylene glycol) diamine. The water (1L) surrounding the dialysis tube was changed once a day during the process for one week.

Conjugation of biotin on FNP -

Fluorescent nanoparticle (FNP -) (2.19 mg, 30.3 µmol) was dissolved in distilled water (4 mL). The total number of acrylic acid units present on the surface of the particles was taken as 1 equiv. Poly(ethylene glycol) 2-aminoethyl ether biotin (69.9 mg, 30.3 µmol) was diluted in 0.5 mL ethanol and then the solution was added into the particle suspension and mixed up to four degree (4°C). About 10 minutes later, a mixture of EDC.HCl {1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride } (23.2 mg, 121.5 µmol) in water was added in the previous solution and was left to stir at 4°C for 24h and then transferred into a cellulose ester dialysis membrane (MWCO: 12-14kDa, Spectrapor) and dialyzed against water to remove the excess of biotin. The water (1L) surrounding the dialysis tube was changed once a day during the process for one week (Scheme 7.3).

Conjugation of antibody on FNP -

Fluorescent nanoparticle (FNP -) was dissolved in distilled water (2 mL). The total number of acrylic acid units present on the surface of the particles was taken as 1 equiv. a mixture 200µL of EDC.HCl {1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride} (4 equiv.) in PBS was added in the previous solution, left for 20mins. Anti-E.coli antibody ab13627 (IgG, abcam) (4 equiv.) was added in and mixed inside the solution. The reaction was left to stir at 4°C for 24h. And then 230µL glycine was added to quench the unreacted sites. The mixture was then transferred into a cellulose ester dialysis membrane (MWCO: 300kDa, Spectrapor) and dialyzed against water to remove excess particles for one night (Scheme 7.4). Scheme 7.4 Synthetic pathway for antibody conjugation to FNP -.

Synthesis of fluorescent polymer chains (FPCs)

Synthesis of negative green fluorescent polymer chains (GFPC -)

BODIPY derivate: BODIPY methacrylate-1 (BDPMA1) was synthesized as described elsewhere. [START_REF] Grazon | Élaboration de nanoparticules fluorescentes à base de BODIPY par polymérisation RAFT en miniémulsion : synthèse, caractérisation et fonctionnalisation de surface[END_REF] Green negative fluorescent polymer chain P(AA 9 -co-APEG 9 -co-BDPMA1 2 ) 20 -TTC-C 12 was synthesized in 1.4-dioxane at 80°C under nitrogen atmosphere. APEG (0.88 mmol, 398 mg), AA (0.93 mmol, 67 mg), TTCA RAFT agent (0.1 mmol, 36.4 mg), DMF (0.54 mmol, 39.7 mg), BDPMA1 (0.24 mmol, 113 mg) were dissolved in 0.97 mL of 1,4-dioxane at room temperature. Then, 0.029 mL of a 0.3 M solution of ACPA in 1.4-dioxane was added. The mixture solution was then purged with nitrogen for 30 min in an ice bath. After this, to initiate the polymerization, the mixture was placed in an oil bath and heated to 80°C. The reaction lasted for 90 minutes and stopped by putting the flask in ice water (Scheme 7.5). Scheme 7.5 Synthetic pathway to get negative green fluorescent polymer chain (GFPC -).

After precipitation in n-pentane, negatively charged green fluorescent polymer chains (P(AA 9 -co-APEG 9 -co-BDPMA1 2 ) 20 -TTC-C 12 ) are obtained with 2 BODIPY fluorophores per chain.

Synthesis of neutral green fluorescent polymer chain (GFPC 0 )

Green neutral fluorescent polymer chain P(APEG 17 -co-BDPMA1 3 ) 20 -TTC-C 12 was synthesized in 1,4-dioxane at 80°C under nitrogen atmosphere. In a typical experiment, APEG (1.77 mmol, 802.2 mg), TTCA RAFT agent (0.11 mmol, 39.4 mg), DMF (0.88 mmol, 0.62 mg), BDPMA1 (0.3 mmol, 140.5 mg) were dissolved in 1.42 mL of 1,4-dioxane at room temperature. Then, 0.029 mL of a 0.3 M solution of ACPA in 1.4-dioxane was added. The mixture was purged with nitrogen for 30min in an ice bath, then placed in an oil bath thermostated at 80°C to initiate the polymerization. After 90 min, the reaction is stopped by immersion of the flask in iced water (Scheme 7.6).

Synthesis of fluorescent pH-sensitive nanoparticles (FANPs)

Synthesis of macroRAFT agents

The PEO 45 -b-PAA 19 -TTC macroRAFT agents were synthesized as described elsewhere 180 .

The P(PEOA 0.5 -co-AA 0.5 ) 22 -TTC macroRAFT agent with equal amounts of AA and PEOA was synthesized in 1,4-dioxane at 75°C under argon atmosphere according to Boisse et al.'s procedure 216 . In the experiment, the RAFT agent TTCA (0.29 g, 0.8 mmol), AA (720 mg, 10 mmol) and PEOA (4.54 g, 10 mmol) were dissolved at room temperature in a flask with 9.9 mL of 1,4-dioxane. 0.1 mL of ACPA solution in 1, 4-dioxane (0.5 M) was then added. The mixed solution was purged for 30 minutes with argon in an ice bath. After this, to initiate the polymerization, the mixture was placed in an oil bath and heated to 75°C. The reaction lasted for 80 minutes and stopped by putting the flask in ice water. The individual molar conversion of each monomer was determined by 1 H NMR in CDCl 3 using DMF as an external standard. The conversion of both monomer was found to be 86%, thus the composition of this copolymer was found to be P(AA 0.5 -co-PEOA 0.5 ) 22 -TTC. The copolymer was precipitated twice in cold diethyl ether in ice water and dried under vacuum at room temperature for 24h.

Synthesis of Polystyrene Nanoparticles (PSNPs)

RAFT polymerization of styrene was performed in the presence of PEO 45 -b-PAA 19 -TTC macroRAFT agents in a one-pot phase inversion process similar to what was reported recently 180 . RAFT polymerization of styrene in the presence of P(PEOA 0.5 -co-AA 0.5 ) 22 -TTC copolymers as macroRAFT agent was performed by the same procedure. In a typical experiment (Scheme7.9), P(PEOA-co-AA)-TTC macroRAFT agent (197mg, 3.2 × 10 -5 mol), styrene (650mg,6.3 × 10 -3 mol) and AIBN (2mg, 1.2 × 10 -5 mol) were dissolved in a flask. The mixed solution was then purged for 30 minutes with argon in an ice bath. After this, to initiate the polymerization, the mixture was placed in an oil bath and heated to 80°C. The reaction lasted for 150 minutes and stopped by putting the flask in ice water. 5 mL of basic water (pH=12.5) were added into the mixture. This biphasic mixture was then placed in an ice bath and an ultrasonic horn (Bandelin electronics, Sonoplus HD 2200) was then placed inside and powered at 130W for 10 minutes. After ultrasonication, the miniemulsion was purged again for 30 minutes in an ice bath with argon. To re-initiate the polymerization, the mixture was placed in an oil bath and heated to 80°C. By this procedure two different PSNPs were obtained with a core-shell structure, while the shell of PSNP1 is PEO 45 -b-PAA 19 , the shell of PSNP2 is P(PEOA 0.5 -co-AA 0.5 ) 22 . Both PSNP1 and PSNP2 have a polystyrene core. Scheme7.9 Synthetic scheme employed for the synthesis of the PSNP1.

Functionalization of PSNPs with fluoresceinamine (FA) 27

In a typical experiment, 0.3 mL of PSNPs (0.1 mg/mL) obtained above was diluted in 4 mL of distilled water in the dark. In parallel, depending on the number of the AA units, 0.5 (PSNP1) or 1 (PSNP2) equivalents of FA (3.35 or 6.7 mg) were dissolved in 0.4 mL ethanol and the solution was added to the PSNPs solution. The mixture was then stirred at 4°C. A solution of EDC (4 equiv., 15 mg) in water (1 mL) was then added to the previous mixture.

After two hours, ethanolamine (2 equiv., 2.3 µL) was added in order to quench the reaction.

The mixture was stirred at 4°C overnight in the dark. Finally, the solution was transferred

Hydrodynamic diameter

The average hydrodynamic particle or polymer diameter (named D h ) was determined by dynamic light scattering (DLS) of the diluted aqueous dispersions at 20°C, with a 4 mW He-Ne laser at 633 nm, using a Zetasizer Nano S90 from Malvern. All calculations were performed using the Nano DTS software.

Size exclusion chromatography (SEC)

The number-average molar mass M n , weight-average molar mass Mw, and the molar mass distribution (polydispersity index M w /M n ) were determined by size exclusion chromatography (SEC) using THF as an eluent at a flow rate of 1 mL.min -1 . For analytical purposes, the acidic functions of the block or statistical copolymers were turned into methyl esters. The copolymers were recovered by drying of the aqueous suspensions. After dissolution in a THF/H 2 O mixture and acidification of the medium with a 1 M HCl solution, they were methylated using an excess of trimethyl-silyldiazomethane. After filtration through 0.45 μm pore size membrane, polymers were analyzed at a concentration of 5 mg.mL -1 in THF. The SEC apparatus is equipped with a sample delivery module (Viscotek GPCmax) and two columns thermostated at 40 °C (PLgel Mixed; 7.5 × 300 mm2; bead diameter, 5 μm). Detection was made with a differential refractive index detector (Viscotek VE 3580 RI detector) and an Ultraviolet-Visible (UV-vis) detector (Waters 486 tunable absorbance detector). The Viscotek OmniSEC software (v 4.6.2) was used for data analysis, and the relative M n and M w /M n were calculated with a calibration curve based on polystyrene standards (from Polymer Laboratories).

Critical micelle concentration (CMC)

The critical micelle concentration (CMC) of GFPCs in water was determined at 20°C or in LB medium determined at 37°C by dynamic light scattering (DLS) using a commercial multi-angle Laser Light Scattering AVL CGS-3 (Compact Goniometer System) equipped with an AVL-5003 digital correlator and a He-Ne Laser (632 nm, 22 mW) [START_REF] Nenninger | Size Dependence of Protein Diffusion in the Cytoplasm of Escherichia coli[END_REF] . Stock solutions of GFPCs (2mg/mL in deionized water, milli-Q grade) were prepared and filtered through a 0.45 µm membrane. Solutions of GFPC -and GFPC 0 of concentrations in the range of 0.01 to 2 mg/mL were prepared by dilution with water milli-Q. The scattered light intensity at 90° was collected for these different samples. By plotting the intensity of scattered light as a function of GFPC -or GFPC 0 concentration, the CMC was determined at the intersection of the straight lines.

Titration of FNP -or GFPC -with Methylene blue

Titration experiments were prepared to characterize the process of FNP -or GFPC - quenching with methylene blue. FNP -solution with an initial concentration of 5x10 -4 µM or GFPC -with an initial concentration of 4x10 -3 µM was placed inside a 4.5mL BRAND Polymethacrylate (PMMA) Cuvette. Absorption and emission spectra were measured.

Methylene blue solutions (1eq, 5 eq, 10eq…) were added inside FNP -or GFPC -solution gradually, and emission spectra of FNP -or GFPC -after adding methylene blue were measured each time. When the decrease of fluorescence intensity of FNP -or GFPC -didn't change anymore, the process was supposed to reach its maximum and the titration was stopped.

Surface Plasmon Resonance (SPR) experiment

Streptavidin chips were prepared on a glass prism (SF 10 with a high refractive index: n 1.707987 at  =830 nm) activated by Reactive Ion Etching (RIE) prior to the thermal evaporation of a 50 nm gold layer. First, a fresh gold layer was immersed in a solution containing 0.1 mM of 1-undecanethiol substituted with a hydroxyl-terminated tetra(ethylene glycol) (EG4-OH) diluted in ethanol over night at room temperature. Before streptavidin deposition, the prisms were thoroughly rinsed three times in pure ethanol for 20 min. The streptavidin solution was spotted on the freshly pre-treated prism surface at different concentrations (10 µM, 1 µM in phosphate buffer (0.4 M) at pH 7.4) and incubated for two hours. The prism was then directly inserted into the SPRi apparatus and the PBS buffer was immediately flowed across the surface at 25 µL/minutes. 1 nM FNP -solution was flowed across the SPRi surfaces containing the spotted streptavidin first. Different images before and after the injection were taken. Then, the surface was washed by PBS buffer. After washing, 1 nM FNP --Biotin solution was flowed across the SPRi surfaces containing the spotted streptavidin, different images before and after the injection were taken, followed by another washing process.

pH measurements

pH measurements were carried by using a glass electrode connected to a PHM210 Standard pH meter from Meterlab. The fluorescence intensity of FANPs was measured every 6 minutes by using a F485/14 filter for excitation and a F535/40 filter for emission. The experiments lasted overnight for ten hours.

Toxicity assessments of fluorescent nanomaterials on E.coli

Bacteria growth medium

Luria broth (LB) medium is a nutritionally rich medium, which is primarily used for the growth of bacteria. M9 Minimal Medium is a microbial growth medium used for the culture of E. coli. This buffered minimal microbial medium contains only salts, so it is traditionally supplemented with glucose, amino acids and vitamins as needed. 2.6x10 -2 µM, 3.9x10 -2 µM and 5.2x10 -2 µM. Di(ethylene glycol) is added to reach 5% of the total volume. Two repeats of the same concentration were carried out during the experiments. The samples were incubated at 37°C in the dark for one hour. After the incubation, the cells were gently centrifuged at 4000rpm for 3 minutes at 4°C and washed with filtered PBS twice to remove free FNPs. The cells were then fixed by adding a fixing solution (4% PFA, 0.03% glutaraldehyde in 1x PBS) for 20 minutes. After the fixing process, cells were washed with PBS for three times and re-suspended in PBS before flow cytometry measurements and bioimaging experiments.

Electroporation to introduce FNPs in E.coli Bacteria

Electroporation is a mechanical method used to introduce molecules into a host cell through the cell membrane. In this procedure, a large electric pulse temporarily disturbs the phospholipid bilayer, allowing molecules like DNA to pass into the cell. Electrocompetent cells are prepared by incubation using 10 % glycerol cold medium. Cells are resuspended in GYT (10 % glycerol, 0.125 % yeast extract, 0.25 % tryptone) cold medium.

An electroporator (Eppendorf, E2510) is used in our experiments. To a Bio-Rad 0.4 cm electrode gap cuvette, we add 50µL electrocompetent cells (8x10 7 bacteria/µL) mixed with different volumes of FNPs to reach different final concentrations of 2.7x10 -4 µM, 1.4x10 -3 µM, 2.7x10 -3 µM, 1.4x10 -2 µM, 2.7x10 -2 µM and 5.2x10 -2 µM. This mixture was placed on ice for 10 minutes before electroporation at the specified voltage (15kV/cm). Immediately after electroporation, 1 mL SOC media (Super Optimal broth with Catabolite repression, a nutrient-rich bacterial growth medium) was added. The cells were then incubated at 37°C for one hour. After the incubation, the cells were gently centrifuged at 4000rpm for 3 minutes at 4°C and washed with filtered PBS twice to remove free FNPs. The cells were then fixed by adding a fixing solution (4% PFA, 0.03% glutaraldehyde in 1x PBS) for 15 minutes. After the fixing process, cells were washed with PBS three times and re-suspended in PBS before flow cytometry measurements and bioimaging experiments.

7.6 Techniques of "Sandwich system" and "Nanoparticle-antibody system" formation 7.6.1 Process of "sandwich system" formation Live E.coli bacterial cells (K-12 BW25113) were used for the following experiments.

Bacterial cultures were prepared overnight from stock cultures inoculated in Luria Broth (LB) growth medium. 1mL of LB growth medium with an initial concentration of 5x10 7 CFU/mL (OD=0.5) of E.coli bacteria was placed inside a 2mL Eppendorf Safe-Lock Tube. The cells were gently centrifuged at 4000rpm for 3 minutes at 4°C and washed with 1mL filtered PBS once and concentrated in 90 µL PBS. 10µL primary antibody (0.4mg/mL) were added into bacterial solution and mixed up at room temperature for two hours incubation. Then the mixture was washed by 400 µL PBS twice and re-suspended in 20 µL PBS. 1 µL streptavidin (0.1 mg/mL) was added to the previous mixture and mixed up at room temperature for 30 minutes, and then washed by 400 µL PBS, twice, and re-suspended in 10 µL PBS. At last, 5 µL FNP --biotin (36 nM) were added to the mixture and mixed up at room temperature for 30 minutes, and then washed by 400 µL PBS twice and then re-suspended in 100 µL PBS before bioimaging experiments.

Secondary fluorescent antibody labelling

Anti-Rabbit IgG (H+L) F(ab') 2 Fragment was conjugated to Alexa Fluor ® 488 fluorescent dye under optimal conditions. 100 µL of LB growth medium with an initial concentration of 5x10 7 CFU/mL (OD=0.5) of E.coli bacteria was placed inside a 1.5 mL Eppendorf Safe-Lock Tube. The cells were gently centrifuged at 4000rpm for 3 minutes at 4°C and washed with 400 µL filtered PBS once and concentrated in 9 µL PBS. 1 µL primary antibody (0.4mg/mL) was added into bacterial solution and mixed up at room temperature for two hours incubation. After, the mixture was washed by 400 µL PBS twice and re-suspended in 9 µL PBS.

1 µL secondary antibody (0.2mg/mL) was added into the solution and mixed up at room temperature for one hour incubation. The mixture was then washed by 400 µL PBS and resuspended in 10 µL PBS before bioimaging experiments.

Process of "Nanoparticle-antibody system" formation

Live E.coli bacterial cells (K-12 BW25113) were used for the following experiments.

Bacterial cultures were prepared overnight from stock cultures inoculated in Luria Broth (LB) growth medium. 25µL of M9 medium with an initial concentration of 1x10 6 CFU/mL (OD=0.4) of E.coli bacteria was placed inside a 0.5mL Eppendorf Safe-Lock Tube. The cells were mixed with FNP-antibody (6.6x10 -3 µM) gently and incubated at room temperature (shaking for one hour). After incubation, bacteria were centrifuged at 4000rpm for 3 minutes at 4°C and washed with 400µL filtered PBS Tween 20 (a wash buffer in immunolabeling techniques. 1mL of LB growth medium with an initial concentration of 5x10 7 CFU/mL (OD=0.5) of E.coli bacteria was placed inside a 2mL Eppendorf Safe-Lock Tube. Different volumes of GFPC -and GFPC 0 solutions were added to this E.coli suspension to reach different final concentrations of 6.6x10 -4 M, 6.6x10 -5 M, 6.6x10 -6 M. Two repeats of the same concentration were carried out during the experiments. The concentration of GFPCs (0.002-0.3mg/mL) in all samples is below the CMC of GFPCs at 37°C (0.4-0.5mg/mL). GFPCs are not likely to form micelles when incubated with E.coli bacteria.

The samples were incubated at 37°C in the dark for one hour. After the incubation, the cells were centrifuged at 4000rpm for 3 minutes at 4°C and washed with filtered PBS twice to remove free GFPCs. The cells were then fixed by adding a fixing solution (4% PFA, 0.03% glutaraldehyde in 1x PBS) for 15 minutes. After the fixing process, cells were washed with PBS for three times and re-suspended in PBS before flow cytometry measurements and bioimaging experiments. The samples were incubated at 37°C in the dark for one hour. After the incubation, the cells were gently centrifuged at 4000rpm for 3 minutes at 4°C and washed with filtered PBS twice to remove free RFPCs. The cells were then fixed by adding a fixing solution (4% PFA, 0.03% glutaraldehyde in 1x PBS) for 15 minutes. After the fixing process, cells were washed with PBS for three times and re-suspended in PBS before bioimaging experiments. The samples were incubated at 37°C in the dark. Every 30 minutes the growth of the cells was monitored using a spectrophotometer, Varian Cary 5000 (Palo Alto, CA USA) that reads the optical density (OD@600nm) of the samples compared to a blank of Modified M9 minimal medium. The pH of the samples was measured using a glass electrode. Emission and excitation spectra of the samples were measured on a SPEX Fluoromax-3 (Horiba Jobin-Yvon).

High-through put screening of bacterial growth inhibition by antibiotics

The assay is performed in a 96 well Falcon Polystyrene Flat Bottom Plate as described above. 150µL of a bacterial solution (1:1000 dilution of an overnight culture) were placed in each well. FANP1 and FANP2 solutions were added into the E.coli solution to reach final concentrations of 8.2x10 -8 M. Three different antibiotics (Chloramphenicol, Kanamycin, and Ampicillin) were added into the bacterial solution to reach an increasing concentrations of 0.5, 1, 5, 10, 50 µg/mL, respectively. Each plate contained two repeats of the same concentration. Controls without antibiotics and one blank with only the Modified M9 minimal growth medium were also prepared.

Preparation of pH sensitive agarose culture plates

Agarose (2% final) was dissolved in 5mL of water and heated at approximately 75°C.

Once the mixture became transparent and cooled down to around 45°C, 5 ml of a 2X solution of the modified M9 minimal medium and different volumes of either FANP1 or with ethanol at different concentrations (70%, 90% and 100%), each concentration was repeated twice and each substitution lasted 10 minutes. After the substitution, a critical point dryer was used to replace ethanol in an ethanol dehydrated specimen with liquid carbon dioxide. Then the liquid carbon dioxide is heated so that at a certain temperature and sample chamber pressure (the critical point) the carbon dioxide passes from the liquid state to the dry gaseous state with little or no damage due to surface tension. Finally, a thin heavy metal platinum layer is applied to the specimen surface by coating in order to increase contrast. Yeast 16, 1421-1427 (2000).

Equations

I would like to thank all the past and present members of the PPSM and LBPA: Andrée I want to express my great thanks to my boyfriend, Xiao, who is always there for me. I cannot imagine how life will be without you. Finally, my greatest love and support is my family. I would like to thank my parents for their continuous support and encouragement.

1

 1 What are bacteria? Why to study them? Bacteria are prokaryotic microorganisms. They are small (few micrometers) single cell microorganisms. They normally exist together in millions. Three different typical shapes of bacteria are encountered: spheres, rods and spirals (Figure 1.1A). Spherical bacteria are usually the simplest ones, called coccus. Rod shaped bacteria are known as bacillus. Spiral bacteria are known as spirillus. Unlike eukaryotic cells, most bacterial cells do not contain intracellular compartments. The study of the diffusion of proteins of different sizes inside a

Figure 1 . 1 A

 11 Figure 1.1 A: Main shapes of bacteria. B: Bacteria number distribution on earth (figure from 4 ).

Figure 1 . 2

 12 Figure 1.2 Structure of E.coli bacteria (figure from 21 )

Figure 1 . 3

 13 Figure 1.3 Perrin-Jablonski diagram (figure from 22 ), IC : internal conversion, ISC : intersystem crossing, S i : singlet electronic state, T i : triple electronic state.

Figure 1 . 4

 14 Figure 1.4 Stokes shift (figure from 24 ).

. 5 )

 5 . Quenching can occur by different mechanisms. Collisional quenching occurs when the fluorophore in the excited state is deactivated by contact with some other molecule which is called the quencher (Figure 1.5A). If fluorophores form non fluorescent complexes with quenchers in the ground state and do not rely on diffusion or molecular collisions the quenching is referred as static (Figure 1.5B).

Figure 1 . 5

 15 Figure 1.5 Schemes of different types of fluorescence quenching (A: Collisional quenching, B: Static quenching, C: Förster resonance energy transfer (FRET)).

Figure 1 . 6

 16 Figure 1.6 Scale bar of the most common nanomaterials compared to that of the main biological structures (figure adapted from 37-40 ).

Figure 1 . 7

 17 Figure 1.7 Structure and spectra of fluorescein in buffer at pH 7: absorption (full line), fluorescence emission (dotted line)(data from life technologies 41 ). Fluoresceinamine is a common amine-reactive derivative of fluorescein. It consists of an amino group which is capable of grafting on carboxylic groups. During my research projects, fluoresceinamine is used as a fluorescent pH indicator which is grafted on polystyrene nanoparticles. I carried out investigation to test if this fluoresceinamine based fluorescent pH-sensitive nanoparticles (FANPs) can be used to detect the growth of E.coli bacteria and their biocompatibility (Chapter 5).

Figure 1 . 8

 18 Figure 1.8 Structure and spectra of Rhodamine 123 in methanol: absorption (full line), fluorescence emission (dotted line)(data from life technologies 50 ).

Figure 1 . 9

 19 Figure 1.9 Emission spectra for the Alexa Fluor dye series (figure from 53 ).

Figure 1 . 10

 110 Figure 1.10 Core structure of the BODIPY fluorophore (inserted) and normalized fluorescence emission spectra of 1) BODIPY FL, 2) BODIPY R6G, 3) BODIPY TMR, 4) BODIPY 581/591, 5) BODIPY TR, 6) BODIPY 630/650 and 7) BODIPY 650/665 fluorophores in methanol(figure adapted from 53 ).

Figure 1 . 11

 111 Figure 1.11 Structures and fluorescence emissions of BDPMA1 (in toluene,  exc =495nm) and BDPMA2 (in ethanol,  exc =540nm) monomers.

charged 59 .

 59 Methylene blue is easily reduced to a colorless form in viable cells leaving them unstained. However, dead cells are unable to reduce methylene blue. Thus stained cells are blue and dead. During my research projects, methylene blue is used to interact with bacteria and fluorescent particles and polymer chains (Chapter 2).

Figure 1 . 12 Figure1. 13

 11213 Figure 1.12 Structure and spectra of oxidized methylene blue in water: absorption (full line), fluorescence emission (dotted line, λ exc =660nm).

Figure 1 . 14

 114 Figure 1.14 Structure and spectra of PKH26 in PBS buffer: absorption (full line), fluorescence emission (dotted line, λ exc =500nm). Propidium iodide is a popular red fluorescent dye. Since propidium iodide is not permeant to live cells, it is commonly used to detect dead cells whose membrane has destabilized and become porous. It has a maximum absorption at 238nm and a maximum emission at 317nm when bound to DNA 64 (Figure 1.15). During my research projects, propidium iodide was used to detect live/dead E.coli bacteria by flow cytometry.

Figure 1 . 15

 115 Figure 1.15 Structure and spectra of propidium iodide bound to DNA: absorption (full line), fluorescence emission (dotted line) (data from life technologies 64 ).

(Figure 1 .Figure1. 16

 116 Figure1.16 Absorption and emission spectra of Aequorea Victoria GFP (figure from[START_REF] Day | The fluorescent protein palette: tools for cellular imaging[END_REF] ).

Figure 1 .

 1 Figure 1.17 timeline of major achievements in the fluorescent proteins field (figure from 65 ).

Figure 1 . 18

 118 Figure 1.18 Different areas of applications for fluorescent proteins in life science (figure from 65 ).

99 (

 99 (Figure 1.19). Surface functionalization or bioconjugation is required to make the fluorescent nanomaterials more biocompatible and able to label biological targets while at the same time preserving their original properties Figure 1.20). In order to increase the water solubility of fluorescent nanomaterials, different coating techniques can be applied, such as hydrophilic thiol coating, silica coating, amphiphilic polymer coating and coating nanoparticles with polyethylene glycol (PEG) (Figure 1.20). Fluorescent nanomaterials can be bioconjugated with biological ligands (e.g. streptavidin, antibody, peptides) in order to make them more biocompatible and also to target biomolecules in cell, bacteria, tissue and living systems (Figure 1.20).

Figure 1 . 19

 119 Figure 1.19 General steps to design functional fluorescent nanomaterials for biosensing and bioimaging.

Figure 1 . 20

 120 Figure 1.20 General strategies to make surface functionalization of fluorescent nanoparticles (figure from 98,100 ) 1.3.3.1 Quantum dots

Figure 1 .

 1 Figure 1.21 Schematic structure of a quantum dot (figure from 101 ).

  spectrum to longer wavelengths. The bigger the QDs, the more red shifted the QDs is 105 . By synthesizing QDs of different sizes, scientists can obtain required emission wavelength. QDs' luminescence covers almost the entire visible spectrum extending to the near infrared range (Figure 1.22).

Figure 1 . 22

 122 Figure 1.22 Increasing the size of QDs resulted in a redshift in the fluorescence emission (figure from 105 )

Figure 1 . 23

 123 Figure 1.23 Bioanalytical applications of bio conjugated QDs (figure from 117 ).

Figure 1 .

 1 Figure 1.24 Structured illumination microscopy (SIM) images of Shewanella oneidensis bacteria with associated amino-PEG QDs, each square has side length 0.42 µm (figure from 122 ).

Figure 1 . 25 Figure 1 Figure 1 . 26

 1251126 Figure 1.25 Fluorescence emission spectra and photographs of organic dye-incorporated silica nanoparticle(50nm) in water (figure from 129 ).Silica nanoparticles have attracted great interests due to their unique properties such as abundant, nontoxic, optical transparency. Furthermore, the ability to capture a large amount of fluorescent dye molecules together in one particle resulted in high brightness, and the silica vehicle can protect fluorophores from molecular oxygen resulting in improved stability. At the same time, the surface of silica nanoparticles can be efficiently conjugated with different biomolecules for biological analytical purpose.

Figure 1 .

 1 Figure 1.27 multi-shell architecture of silica nanoparticle applications (figure from 141 ).

  attract great attention for fluorescent labelling and sensing technologies because of their high brightness, high photostability, abundant surface modification and easily bioconjugation, and most importantly low toxicity for living systems. During my research project I found that all the fluorescent organic nano-objects used for fluorescent labeling and sensing are nontoxic for bacterial growth and can be used for live bacterial imaging applications. The principle of design for any fluorescent organic polymer nanoparticles is the same: gathering as many fluorohores as possible together in one nano-object. Unlike single organic dye such as FITC or rhodamine, each organic nano-object can contain thousands of embedded fluorophores that are encapsulated in a polymer shell and protected from the outside environment. Because of the high number (up to thousands) of dye molecules incorporated into one single particle, organic polymer particles are much brighter than individual organic molecules. At the same time, since they are encapsulated, fluorophore molecules are protected from outside environment and less affected by the external environment and are more stable (less photobleaching or blinking as well) 150 . Bioconjugation of various biological functional groups such as biotin, avidin, antibody, antigenic peptides, to the surface of polymer nanoparticles allow for their labeling of cells, tissues, organisms, bacteria and other living systems (Figure 1.28).

Figure 1 . 28

 128 Figure 1.28 Organic polymer nanoparticles with multifunctional bioconjugations (figure adapted from 151 ).

Figure 1 . 29

 129 Figure 1.29 Preparation of polymer nanoparticles by the miniemulsion method (figure from 155 ).

Figure 1 . 30

 130 Figure 1.30 Preparation of polymer nanoparticles by the reprecipitation method.

Figure 1 . 31

 131 Figure 1.31 Schematic preparation of polymer nanoparticles by the self-assembly method.

Figure 1 . 32

 132 Figure 1.32 Schematic preparation of polymer nanoparticles by direct polymerization (figure adapted from 160 ).

  demonstrated that conjugated polymer nanoparticles generated by using an amine containing poly(p-phenylene ethynylene) (PPE) can be used to perform live cell imaging. Different cells including baby-hamster kidney (BHK) and BALB/C 3T3 (mouse embryonic fibroblast) were incubated with conjugated polymer nanoparticles for different times varying from one hour to several days in culture media to study the cell permeability, cellular toxicity and photostability of conjugated polymer nanoparticles. Microscopy images of live BALB/C 3T3 (a) and fixed 3T3 cells (b) stained by the conjugated polymer nanoparticles overnight are shown in Figure 1.33. The images show that the conjugated polymer nanoparticles were cell permeable without any inhibition of cell viability and can accumulate particularly in the cytosol. Furthermore, compared with commercial dyes, conjugated polymer nanoparticles possess high resistance to photobleaching 162 .

Figure 1 . 33

 133 Figure 1.33 Fluorescence images of live (a) and fixed (b) cells. (a) BALB/C3T3 cells were incubated conjugated polymer nanoparticles (green) and Hoechst dye (blue). (b) Fixed BALB/C 3T3 cells for confocal microscopy 162 .

Figure 1 .

 1 Figure 1.34 (a) Fluorescence microscopy images of coating conjugated polymer nanoparticle on E. coli to form microparticles (b) SEM images of E. coli alone and E. coli microparticles (figure from 163 ).

  developed a drugencapsulated, pH-responsive, surface charge-switchable PLGA-PLH-PEG (poly(D,L-lactic-coglycolic acid)-b-poly(L-histidine)-b-poly-(ethylene glycol)) nanoparticles system for bacterial infections therapy (Figure 1.35). These nanoparticle drug carriers are negatively charged and they are designed to switch their surface charge to positive and bind strongly to the bacterial membrane which is negatively charged in an acidic environment and deliver drugs to bacteria with decreasing pH. These PLGA-PLH-PEG NPs are considered to be the first step for developing drug carriers systemically which can target and potentially treat bacterial infections associated with acidic activity 167 .

Figure 1 .

 1 Figure 1.35 Drug-encapsulated, pH-responsive, surface charge-switchable PLGA-PLH-PEG nanoparticles system for bacterial infections therapy (figure from 167 ).

Figure 1 .

 1 36. David H. Gracias and his team 171 developed a two-antibody-based technique to attach and release bacteria to polystyrene beads (around 500nm) to form bacteria cargo from patterned surfaces. Polystyrene beads were coated with FITC-labeled bacteria-capturing antibody and were bound to a second antibody which is stabilized on gold patterns. These PS beads can be used to capture E.coli bacteria (Figure 1.37 left). The bacteria-nanoparticle cargo can be released on-demand from the gold patterns with imidazole or EDTA, and these motile bacteria are observed to propel their cargo swimming in growth media (Figure 1.37 right).

Figure 1 . 36 Figure 1 .

 1361 Figure 1.36 Streptavidin conjugated nanoparticle to form bacteria-nanoparticles cargo (figure from 174 ).

  incorporated in agarose gel to form sensor film. The films were then used for real time monitoring for bacterial growth and exhibited a visible color change from blue to green as shown in Figure 1.38.

Figure 1 .

 1 Figure 1.38 Representation of the mechanism for monitoring bacterial growth by polyaniline nanoparticles based colorimetric sensor (figure from 175 ).

1. 4

 4 Novel fluorescent nanomaterials for bioimaging and biosensing: application on E.coli bacteriaIn the quest for very bright and stable labels, we envisioned the design and characterization of novel polymer-based, self-stabilized, fluorescent nanoparticles (FNPs, 60nm) and fluorescent polymer chains (FPCs, 5nm). The main structure of these fluorescent organic nanomaterials are shown in Figure1.39 f, g, i, j. They are compared with single organic dyes which were used to form these nanomaterials (Figure1.39 a, b, c) and common fluorescent labels which have been described in last section such as: GFP, QDs, dye doped silica nanoparticles (Figure1.39 e, d, h). I have used these nanomaterials to study the internal and external bioimaging and biosensing application on E.coli bacteria, the strategies are generally shown in Figure1.40.

Figure 1 .

 1 Figure 1.39 Main structures of fluorescent organic nanomaterials and their comparison. a) BDPMA1 (BODIPY methacrylate-1), b) BDPMA2 (BODIPY methylnaphthalene-2), c) FA (Fluoresceinamine 27,177 ), d) QDs (Quantum dots 28,178 ), e) GFP (Green fluorescent proteins 179 ), f) GFPC -(negative Green fluorescent polymer chains), g) RFPC -(negative Red fluorescent polymer chains), h) Silica nanoparticle 141 , i) FANP1 (fluoresceinamine based fluorescent pH-sensitive nanoparticles), j) FNP - (negative Fluorescent nanoparticles).

Figure 1 .

 1 Figure 1.40 Scheme of the envisioned fluorescent nanomaterials for bioimaging and biosensing: application on E.coli bacteria. (A) FNPs: Fluorescent nanoparticles. (B) FNP-bioconjugation. (C) GFPCs: Green fluorescent polymer chains. (D) RFPCs: Red fluorescent polymer chains. (E) FANPs: Fluorescent pH-sensitive nanoparticles

  order to investigate the biological applications of FNPs on E.coli bacteria, different strategies have been developed as shown in Figure 1.40. One strategy is to internalize fluorescent nanomaterials (Figure 1.40A: FNPs, C: GFPCs, D: RFPCs) in E.coli bacteria, and investigate which part of bacteria is labeled (or not). The second idea is that we can conjugate biomolecules on the surface of the particles to target the bacterial membrane (Figure 1.40B:

Figure 1 .

 1 40A) into E.coli bacteria. To control if the FNPs are indeed internalized, I developed a protocol based upon FNPs luminescence quenching by methylene blue (see Chapter 2).

Figure 1 .Figure 1 .

 11 Figure 1.40E) are used to measure the presence of E.coli bacteria growth. I will show how bacteria influence the fluorescence intensity of FANPs after incubating E.coli bacteria with pH sensitive FANPs. I have demonstrated that these FANPs can be used for a rapid, accurate, reproducible and highly sensitive detection of E.coli bacterial growth. I will show that such FANPs can be used for continuous monitoring for real-time detection in long time scales and can be used for screening of a large number of samples for high-throughput applications (see Chapter 5).

Figure 2 . 1 Figure 2 . 2

 2122 Figure 2.1 BDPMA1 monomer in toluene (black lines) and FNP -(grey lines) in water (pH=7.0): absorption (A, B full lines), fluorescence emission (A dotted lines, λ exc =495nm) and excitation (B dotted lines, λ emi =543nm) normalized spectra.

2 . 2 .Figure 2 . 3

 2223 Figure 2.3 FNP -(black lines), FNP + (dark grey lines), FNP-PEG + (light grey lines) in water (pH=7.0): absorption (A, B full lines), fluorescence emission (A dotted lines, λ exc =495nm) and excitation (B dotted lines, λ emi =543nm) normalized spectra.

Figure 2 . 4

 24 Figure 2.4 Growth curves of E.coli bacteria alone (red lines) and E.coli bacteria incubated with FNP - (yellow lines), FNP + (blue lines) and FNP-PEG + (purple lines) at 6.6x10 -3 µM in LB (A) or M9 media(B). The error bars indicate the difference between the three replicates in the experiment (<9%).

  5) in the LB growth medium (rich medium, §7.4.1) was placed inside an Eppendorf Safe-Lock Tube in the presence of FNP -(at 3 different concentrations-2.6x10 -2 µM, 3.9x10 -2 µM, 5.2x10 -2 µM) and 5% DEG. The samples were incubated at 37°C in the dark shaking for one hour and washed twice with PBS to remove free FNP -. The cells were then fixed by adding a fixing solution (4% PFA, 0.03% glutaraldehyde in 1x PBS), then washed three times with PBS and finally re-suspended in PBS before flow cytometry measurements and bioimaging experiments. Accurate measurements of fluorescent cells (Figure 2.5) after incubation of E.coli with different concentrations of FNP -(2.6x10 -2 µM, 3.9x10 -2 µM, 5.2x10 -2 µM) were obtained by Flow Cytometry.

Figure 2 . 5 Figure 2 . 6

 2526 Figure 2.5 Percentage of fluorescent cells for E.coli incubation with FNP -after DEG (5%) treatments.E.coli interacting with FNP -after chemical treatment was observed by epifluorescence microscopy. In Figure2.6 one can see that some cells are associated with the FNP -and several FNPs are sometimes found on one bacterium. However, the FNPs seemed to be attached to the membrane and not inside the cells.

  Accurate measurements of live, dead, total bacteria and fluorescent cells percentage after E.coli electroporation with different concentrations of FNP -(2.7x10 -4 -5.2x10 -2 µM) are obtained by Flow Cytometry (Chapter 7, §7.8) (Figure 2.7). Live cells have intact membranes and are impermeable to dyes such as propidium iodide (PI-see Chapter 1 §1.3.1), which only leaks into dead cells when the membrane is compromised. After addition of PI, we can measure the percentage of dead cells in each sample.

Figure 2 . 7 Figure 2 . 2 Figure 2 . 8 Figure 2 . 9

 27222829 Figure 2.7 Flow Cytometry of E.coli interacting with FNP -after electroporation, A) competent cells, B) competent cells after electroporation, C) competent cells and FNP -after electroporation (2.7x10 -9 M). X axis: green fluorescence, Y axis: red fluorescence. (R1: live non-fluorescent cells, R2: live green fluorescent cells, R3: dead red-green fluorescent cells, R4: dead red fluorescent cells). As shown in Figure 2.7A, 10% cells (R4) are dead after competent cells preparation while in Figure 2.7B one can see that 50% (R4) of the cells are dead after electroporation. In Figure 2.7C when competent cells are exposed to FNP -and electroporation then 60% of cells are dead and labeled by FNP -(R3), 40% of cells survived, only 8% of cells are alive and labeled by FNPs (R2). When the particle concentration is increased, the percentage of live

Figure 2 .Figure 2 . 10 Figure 2 . 11

 2210211 Figure 2.10 Bioimaging of one E.coli interacting with FNP -after electroporation. (A): phase contrast image, IT=400ms; (B): fluorescence image, IT=5s; (C): overlap of images A and B (60X objective, Particle concentration: 2.7x10 -3 µM).Larger images (Figure2.11) show that after electroporation, many cells are associated with FNP -and several nanoparticles on one bacterium are often observed.

Figure 2 . 12

 212 Figure 2.12 Overlap of phase contrast and fluorescence images of E.coli interacting with FNP + (A, B) and FNP-PEG + (C, D). (60X objective, Particle concentration: 2.7x10 -3 µM). Additional images are shown on FigureS2.3 in Section 2.6.

Figure 2 .Figure 2 . 13

 2213 Figure 2.13 Absorption (full lines) and emission (dotted lines) spectra of FNP -(black) and MB (grey) in water (A); Fluorescence spectra recorded during the titration of FNP -with MB (B,  exc =495nm); Fluorescence spectra recorded of MB in the same condition without FNPs (C,  exc =495nm); Relative decrease of fluorescence intensity of FNP -as a function of the concentration of methylene blue (D).

Figure 2 . 14 (Figure 2 .Figure 2 .

 21422 Figure 2.13C. Since the core of the particle is formed by poly(styrene-co-BDPMA1), the BODIPY fluorophores are in distinct environments. The plot proves our hypothesis that there are two populations of BDPMA1s inside the particle, one on the surface of the core is more accessible (a) to quencher methylene blue and the other one deep in the core is being inaccessible or buried (b). These results corroborate a previous study on the FNP -where a dual population was also postulated to explain their fluorescence decays 4 .

Figure 2 . 15

 215 Figure 2.15 Mechanism of differentiation of particles inside or outside bacteria by addition of methylene blue and selective quenching. Light green refers to a quenched FNP, while dark green refers to a fluorescent FNP (unquenched).

Figure 2 . 15

 215 Figure 2.15 Comparison of electrocompetent E.coli interacting with FNP -before (A, C) and after (B, D) quenching with methylene blue (60X objective, A, B: phase contrast images, IT=200ms; C, D: fluorescence images, IT=5s). Additional images are shown in Figure S2.4 in Section 2.6.

Figure 2 . 16

 216 Figure 2.16 Fresh porcinis (A) and air dried porcinis (B) (adapted from 189 ).

Figure 2 .Figure 2 . 17

 2217 Figure 2.17 Phase diagram of CO 2 (A); Surface of rose blossom after critical point drying (B) and air drying (C) (figure from 189 ).

Figure 2 .Figure 2 .Figure 2 .

 222 Figure 2.18 SEM images of untreated E.coli (A) and electrocompetent cells (B). Additional images are shown in Figure S2.5 in Section 2.6. From the flow cytometry experiments, one can notice that 50% cells were dead after electroporation. And indeed, electroporation causes serious damages to bacteria as shown in Figure 2.19. After electroporation, holes form on the bacteria membrane, especially at the poles of the bacteria which appeared to be more frangible. After electroporation, cells in different states were also observed (Figure 2.19B). After the breakage, some cells released their cytoplasm and are dead.

Figure S2. 1 Figure S2. 2 3

 123 Figure S2.1 Epifluorescence microscopy imaging of E.coli interacting with FNP -after DEG treatment.

Figure S2. 4 Figure S2. 5 Figure S2. 6

 456 Figure S2.4 Comparision of electrocompetent E.coli interacting with FNP -after electroporation and before (A, C)-after (B, D) quenching with methylene blue (60X objective, A, B: phase contrast images(IT=200ms); C, D: fluorescence images(IT=5s)).

Scheme 3 . 1

 31 Scheme 3.1 Hypothesis of particle based "sandwich system"

Scheme 3 . 3

 33 Scheme 3.3 Process of sandwich system formation

Figure 3 . 1

 31 Figure 3.1 FNP -(red lines), FNP --Biotin (orange lines, FNP --biotin-streptavidin (green lines), FNP -biotin-streptavidin-antibody (blue lines) in PBS: absorption (full lines) and fluorescence emission (dotted lines,  exc =495nm) normalized spectra.

Figure 3 . 2

 32 Figure 3.2 DLS signals of FNP -(A), FNP --biotin (B), FNP --biotin-streptavidin(C) and FNP --biotinstreptavidin-antibody (D) in PBS (pH 7).

Figure 3 . 3

 33 Figure 3.3 Streptavidin gold chip for SPR.

Figure 3 .

 3 Figure 3.4A corresponds to the differential image at time t=0 where no FNP -was injected. Figure 3.4B, C, D are images taken during the association step, at a total FNP - concentration of 1 nM. Since there is no biotin conjugation, we observed a non-specific binding between particles and streptavidin. The spots became slowly bright and the contrast increased until it reached a maximum (Figure 3.4D).The kinetic curve of FNP -and streptavidin interaction confirms this observation (Figure3.5A). Moreover, there is a higher relative reflectivity intensity for the spots at 10 µM streptavidin than the one at 1 µM which indicates that increasing the streptavidin concentration favors the non-specific interaction between particles and streptavidin. Immediately following the injection of FNP -, the surface was continuously rinsed with buffer. The background signal returned to its original level while the relative reflectivity intensity in the streptavidin containing spots went back to low level (Figure3.4E). We also observed a huge decrease of relative reflectivity intensity from the kinetic curve after washing for both concentrations of streptavidin (Figure3.5A).

Figure 3 . 4 Figure 3 . 5

 3435 Figure 3.4 SPRi difference images of the biochip surface at different times during the course of the experiment. 2 different streptavidin solution concentrations (10 µM, 1 µM) were spotted as shown in Figure 3.3. A: t = 0 s, no FNP -injected. B, C, and D are images taken at t = 1min, t = 2 min and t = 4 min respectively after 1 nM FNP -injection. E is the image taken 30 min after the injection was stopped and washed by buffer. F, G and H are images taken at t = 30 s, t = 2 min, t = 4 min respectively after 1 nM FNP --biotin injection.

Figure 3 . 4 F

 34 , G, H are images taken during the association step. As we know, streptavidin-biotin bonding is one of the strongest known non-covalent interactions (K d of 10 -14 mol/L)195 . With biotin conjugated particles, there is a very strong non-covalent interaction between particles and streptavidin, the spots became bright quickly and the contrast increased to reach the maximum very quickly (Figure3.4 H).

3 Figure 3 . 6 A 3 Figure 3 . 7

 336337 Figure 3.6 A: Principle of secondary antibody labeling primary antibody on bacteria. B: Absorption and emission spectra of Alexa Fluor® 488 (data from life technologies 196 ).

Figure 3 .

 3 8A is the phase contrast image of bacteria, after formation of sandwich system and particles could label the bacterial membrane as shown by fluorescence in Figure 3.8B.

Figure 3 . 8

 38 Figure 3.8 Bioimaging of particle based sandwich system targeted on bacterial surface. (60X objective, A: phase contrast image, IT=200ms. B: fluorescence image overlap with phase contrast image, IT=5s). Additional images are shown in Figure S3.2 in Section 3.5.

described in Chapter 2 (Figure 3 . 9

 239 Figure 3.9 SEM images of E.coli with FNP --biotin without streptavidin (A, B) and with streptavidin (C, D). Additional images are shown in Figure S3.4 in Section 3.5.

  4, IBIDI) was used as shown in Figure 3.10. 30 µL FNP --biotin particle solutions in PBS at different concentrations (0.01, 0.04, 0.07, 0.1 nM) were injected into the µ-Slide channels. After 10 minutes waiting to let the particles attach on the surface, 1mL PBS was flowed three times to wash away excess FNP -.

Figure 3 . 10 3 Figure 3 . 11

 3103311 Figure 3.10 Flow experiments for FNP --biotin (a: 0.01, b: 0.04, c: 0.07, d: 0.1 nM) on µ-Slide channel (figure adapted from IBIDI 197 ). Images of FNP --biotin particles attached on the slide surface before and after washing are shown in Figure 3.11. After injection of particles inside the channels, FNP --biotin particles stick on the surface (Figure 3.11 A). It is noteworthy that even with the lowest concentration (0.01 nM) of particle solution, the number of FNP --biotin is huge. A lot of particles stick to the surface giving a strong fluorescent background and particles can't be identified. After the first washing (Figure 3.11 B), the background signal became less strong, however particles could still not be identified clearly , the particles surface density seems too high, we aim at less surface coverage. After the second washing (Figure 3.11 C), a lot of particles still stick on the surface and there is no visible difference between the four different concentrations of FNP --biotin used. It means that the lowest concentration (0.01nM) is already enough, so I washed channel "a" (Figure 3.10) which contained the

Figure 3 .Figure 3 . 13 5 .

 33135 Figure 3.13A. It is very difficult to distinguish individual cell or to tell if they are linked to particle from the fluorescence image (Figure 3.13B), so I tried to wash free bacteria away. The cells are gently washed by PBS twice. After the first wash, most of the unbound cells have been washed away (Figure 3.13C). However there are still a lot bacteria remaining.Even if all the bacteria are associated with particles (Figure3.13D), it is still difficult to identify each bacteria and particle location.

Figure 3 .

 3 14A, one can notice that these acrylic acids units are "hidden" by the poly(ethylene oxide) (PEO) which in the most outside layer of the shell. This structure may affect the accessibility of the reaction between the biotin and acrylic acids. Thus, I changed to another type of fluorescent nanoparticle (FNPB, Figure 3.14B) synthesized by Chloé Grazon 27 with a different shell architecture which might be more efficient for antibody conjugation experiments.

Figure 3 . 14

 314 Figure 3.14 Structures of FNP -(A) and FNPB (B)-(figure adapted from 27 ).

Scheme 3 . 4 2 .Figure 3 .

 3423 Scheme 3.4 Synthetic pathway for antibody conjugated on FNPB.As said before, it was difficult to characterize the conjugation by conventional method (e.g. NMR or IR). Absorption and fluorescence spectra of FNPB and FNPB-antibody were recorded as shown in Figure3.15 and their main spectroscopic data are given in Table3.2.The FNPB particle in PBS shows a maximum absorption band at 529nm, and a maximum emission band at 541nm. The FNPB-antibody shows very similar absorption and fluorescence spectra as FNPB (Figure3.15A) except that the FNPB-antibody exhibits a more intense bond in the UV region at around 380nm. This is probably due to the light scattering by the conjugated antibody which are relatively large (150 kDa 194 ). The excitation spectra for FNPB and FNPB-antibody have the same aspect and superimpose to their respective absorption spectra (Figure3.15B). In PBS the fluorescence quantum yield of FNPB is 42% and the one for FNPB-antibody is 28% indicating that formation of the "nanoparticle-antibody" may decrease the quantum yield of FNPB.

Figure 3 . 3 Figure 3 . 15

 33315 Figure 3.15 Bioimaging of "nanoparticle-antibody" targeted on bacterial surface. (100X objective, A: bright field image, IT=500ms, B: fluorescence image overlap with bright field image, IT=500ms). Additional images are shown in Figure S3.5 in Section 3.5.

Figure 3 . 16 Figure 3 . 17

 316317 Figure 3.16 Strategy for particle based "sandwich system"In order to characterize the formation of sandwich, I divided it into four different parts: FNP -, FNP --biotin, FNP --biotin-streptavidin, FNP --biotin-streptavidin-antibody. I analyzed each part of the sandwich, and studied their properties and compared them with each other.

Figure S3. 1

 1 Figure S3.1 Comparison of E.coli -secondary fluorescent antibody (A, B) and E.coli-primary antibodysecondary fluorescent antibody (C, D)(60X objective, A,C: phase contrast image, IT=200ms. B, D: fluorescence image, IT=5s).

Figure S3. 2 3 4 Figure S3. 5

 2345 Figure S3.2 Bioimaging of particle based sandwich system targeted on bacterial surface. (60X objective, A: phase contrast image, IT=200ms. B: fluorescence image overlap with phase contrast image, IT= 5s).

Scheme 4 . 1

 41 Scheme 4.1 Synthetic pathway for negative green fluorescent polymer chain (GFPC -).

2

 2 Scheme 4.2 Synthetic pathways for neutral green fluorescent polymer chain (GFPC 0 )(A), negative red fluorescent polymer chain (RFPC -)(B), positive red fluorescent polymer chain (RFPC -)(C).

Figure 4 . 1 .

 41 The BDPMA1 conversion is determined by SEC using UV-vis detection (λ abs = 528 nm). It shows narrow symmetric peaks that are shifted towards shorter retention volumes (corresponding to higher molecular weight and longer polymer chains) upon progressing polymerization (Figure4.1A, C). For GFPC -, the polymerization reached 60% conversion of BDPMA1 after 4 hours (Figure4.1B)

Figure 4 . 1

 41 Figure 4.1 Normalized size exclusion chromatograms (UV signal, λ abs = 528 nm) in THF for GFPC -(A, B) and GFPC°(C, D), at different times of monomer conversions, and (inset A, C) evolution of the number-average molar mass (M n , PS calibration) and M w /M n vs. monomer conversion, and (inset B, D) evolution of BDPMA1 conversion/% vs. time/minutes. The straight line corresponds to the theoretical M n values (Equation 7.2 in Chapter 7). The indicated percentages % corresponds to the BDPMA1 conversion.

Figure 4 . 2

 42 Figure 4.2 Intensity of scattered light vs. molar concentration of GFPC -(A, B) and GFPC 0 (C, D). The intersection of the straight lines corresponds to the critical micelle concentration (CMC).

Fluorescence

  and absorption spectroscopy analysis of GFPCs are displayed on Figure 4.3 and their main spectroscopic data are given in Table

4 .

 4 3A) 181 . The excitation spectra for GFPCs and the BDPMA1 monomer virtually superimpose with their respective absorption spectra (Figure 4.3B), showing that the absorbing and emitting species are identical. DLS signals of GFPC 0 (142nm) and GFPC -(84nm) are shown in

Figure 4 . 4 ,Figure 4 . 3 Figure 4 . 4

 444344 Figure 4.4, compared with theoretical diameter (4-6nm), the measured diameters are much higher indicating presence of aggregation of GFPCs inside aqueous solutions.

182,Figure 4 . 5

 45 Figure 4.5 BDPMA1 monomer and GFPCs in toluene: absorption (A, B full lines), fluorescence emission (A dotted lines, λ exc =495nm) and excitation (B dotted lines, λ emi =548nm) normalized spectra.

Figure 4 . 6

 46 Figure 4.6 Growth curves of E.coli bacteria alone (black lines) and E.coli bacteria incubated with GFPC -(dark grey lines) and GFPC 0 (light grey lines) at 6.6x10 -5 M in LB medium(A) or M9 medium(B). The error bars indicate the difference between the three replicates in the experiment (<6.6%).

Figure 4 .Figure 4 . 7

 447 Figure 4.7 Percentage of fluorescent cells for GFPC 0 (A), GFPC -(B) total fluorescent cells (dotted lines), live fluorescent cells (full lines). Flow cytometry analysis of E.coli incubation with GFPC 0 (C), GFPC -(D) at the concentration of 6.6x10 -5 M, x axis: green fluorescence, y axis: red fluorescence(R1: live non-

Figure 4 . 8 ).

 48 Compared with GFPC 0 (Figure 4.8B) and GFPC -(Figure 4.8C), it is found out that under the same concentration and bioimaging conditions, the bacteria in Figure 4.8C are brighter than the one in Figure 4.8B. From

Figure 4 . 8

 48 Figure 4.8 Fluorescence image (60X objective, IT=10s) of E.coli alone (A) and E.coli interacting with GFPC 0 (B) and GFPC -(C). Additional images are shown in Figure S4.1 in Section 4.9.

Figure 4 . 9 AFigure 4 . 9

 4949 Figure 4.9 Absorption and emission spectra of GFPC -(λ exc = 495nm) and MB (λ exc =660nm) (A), titration process of MB to GFPC -(B). Percentage decrease of fluorescence intensity of GFPC -as a function of the concentration of methylene blue (C).

Figure 4 .

 4 10A (Equation 7.3 in Chapter 7). A linear Stern-Volmer plot is generally indicative of a single class of fluorophores, all equally accessible to quencher 23 . Since in our case the Stern-Volmer plots deviate from linearity toward the x-axis, it means that two or several fluorophore populations are present, and there are classes are not accessible to quencher. However in GFPC -, only BPDMA1 is the fluorophore, so it can be assumed that there are two or more populations of BDPMA1s, one of which is accessible (a) to quencher methylene blue and the others being inaccessible or buried (b). At the same time, in Figure 4.9C, there are three slopes for the fluorescence intensity decreasing process of GFPC -indicating that there might be three populations of BDPMA1 as we have hypothesized. In this case the Stern-Volmer plot will display a downward curvature which is exactly the one show in Figure 4.10A. In addition, a slight blue shift of the emission of GFPC -is observed in the titration process (Figure 4.9 B), the emission spectrum shifts progressively to shorter wavelengths with increasing MB concentrations. This is also probably due to selective quenching of exposed BDPMA1 versus buried BDPMA1 and indicates that those BDPMA1 emitting at larger wavelengths are quenched more readily than the shorter wavelength BDPMA1. This spectrum shows that the quenched BDPMA1 display an emission maximum at 548 nm. The protected BDPMA1 display an emission maximum at 540 nm. A modified Stern-Volmer plot (Figure 4.10 B) is made according to the modified Stern-Volmer equation (Equation

Figure 4 .

 4 Figure 4.10 Stern-Volmer (A) and modified Stern-Volmer plots (B) for GFPC -quenched by methylene blue.

Figure 4 .Figure 4 . 1 Figure 4 . 12

 441412 Figure 4.11 E.coli interacting with GFPC -after quenching with methylene blue (100X objective, A: bright field image, IT=500ms; B: fluorescence image, IT=5s). Additional images are shown in Figure S4.2 in Section 4.9.4.4.3 Multi-color imaging application of GFPC -

Figure 4 . 14 .

 414 It has two bands of absorbance at 600nm and 646nm and it emits in the far-red / near infra-red (NIR) and has maximum band at 697nm when bound to dsDNA.

Figure 4 . 13

 413 Figure 4.13 Structure and spectra of DRAQ 5 in PBS buffer: absorption (full line), fluorescence emission (dotted line, λ exc =600nm)(figure adapted from 61 ). During our experiments, DRAQ5 was used to stain the bacterial DNA in order to discriminate the location of DNA and GFPC -. DRAQ5 was first tested on E.coli for the ability to label bacterial DNA, DRAQ5 (1:1000 dilution) with an initial concentration of 5mM was added into a bacterial suspension and incubated for one hour. Samples were washed twice with PBS before the microscopy experiments (Figure 4.14).

Figure 4 . 14 9 .

 4149 Figure 4.14 Images of DRAQ5 labeling the DNA of E.coli (60X objective, A: phase contrast image, IT=100ms; B: green fluorescence image, IT= 20s; C: red fluorescence image, IT=20s; D: overlapped image of A and C). Additional images are shown in Figure S4.3 in Section 4.9. After successfully using DRAQ5 to label the DNA of E.coli, DRAQ5 and GFPC -were then used for multi-color labelling of bacterial cytoplasm. DRAQ5 (1:1000 dilution) with an initial concentration of 5mM was added at the same time with GFPC -(6.6x10 -5 M) to E.coli suspension in M9 medium to incubate for one hour. Samples were washed two times with PBS before the microscopy experiments.

Figure 4 . 15 9 . 4 . 4 . 3 . 2

 41594432 Figure 4.15 Bioimaging of E.coli bacteria incubated with GFPC -and DRAQ5 (60X objective A: phase contrast image, IT=100ms; B: green fluorescence image, IT= 20s; C: red fluorescence image, IT=20s). Additional images are shown in Figure S4.4 in Section 4.9.

Figure 4 . 16

 416 Figure 4.16 Structure and spectra of PKH26 in PBS: absorption (full line), fluorescence emission (dotted line, λ exc =500nm).

Figure 4 .

 4 Figure 4.17B, PKH26 could be used to label bacterial membrane successfully which indicatesthat GFPC -can be used to discriminate the membrane and GFPC -location. However the efficiency of PKH26 to label the bacterial membrane is not very high, I measured for seven images and about 10.1% cells are labelled successfully which compared with the efficiency of GFPC -(80%) to label bacterial cytoplasm, it is relatively low.

Figure 4 . 17 9 .

 4179 Figure 4.17 Images of PKH26 labeling the membrane of E.coli (100X objective, A: bright field image, IT=500ms; B: fluorescence image, IT=5s). Additional images are shown in Figure S4.5 in Section 4.9.After successfully using PKH26 to label the membrane of E.coli, PKH26 and GFPC -were then used for multi-color labelling of bacterial cytoplasm (Figure4.18).

Figure 4 . 9 .

 49 Figure 4.18 E.coli interacting with GFPC -and PKH26 (A: bright field image, IT=500ms; B: orange fluorescence image, IT=5s; C: green fluorescence image, IT=5s; D: overlapped image of B+C). Additional images are shown in Figure S4.6 in Section 4.9. The bacterial cytoplasm was targeted by GFPC -(Figure 4.18C) which is the green part, and the bacterial membrane was labelled by PKH26 (Figure 4.18B) which is the yellow part.

  Indeed, positive and negative red fluorescent polymer chains (RFPCs, Scheme 4.2) have been synthesized with similar architecture as GFPCs. The BODIPY methylnaphthalene-2 (BDPMA2) is introduced in the synthesis process, the additional aromatic moieties on the pyrrole of the BODIPY monomer shift fluorescence emission to red by conjugation extension of the  system. These RFPCs are useful to try for biological imaging application and we wish to use RFPCs and GFPCs for multi-color imaging. RFPCs were recently synthesized by Muriel Coquelin and the characterization is still under investigation. The exact number of BDPMA2 per chain and the BDPMA2 conversion rate has not yet been determined. However, I carried out some preliminary spectroscopy measurements to obtain an overall understanding of RFPCs which are necessary for bioimaging experiments. The fluorescence and absorption spectroscopy analysis of RFPCs are displayed on

Figure 4 .

 4 Figure 4.19 and their main spectroscopic data are given in Table 4.4. The BDPMA2 monomer in ethanol shows a red shift compared with the one of BDPMA1 (Figure 4.19A). The maximum of absorption and fluorescence emission are found at 552nm and 594 nm, exhibiting a Stokes shift of 1281cm -1 (42nm). The RFPCs show very similar absorption and fluorescence spectra with a maximum absorption at 552nm (RFPC + ), 553nm (RFPC -).

Figure 4 .Figure 4 .Figure 4 .

 444 Figure 4.19 BDPMA1 monomer in toluene and BDPMA2 monomer in ethanol: absorption (A, B full lines), fluorescence emission (A dotted lines, λ exc BDPMA1 =495nm, λ exc BDPMA2 =540nm) and excitation (B dotted lines, λ emi BDPMA1 =548nm, λ emi BDPMA2 =587nm).A B

Figure 4 .

 4 Figure 4.22 BDPMA2 monomer and RFPCs in toluene: absorption (A, B full lines), fluorescence emission (A dotted lines, λ exc =540nm) and excitation (B dotted lines, λ emi =587nm) normalized spectra.

Figure 4 .Figure 4 . 23

 4423 Figure 4.23 Growth curves of E.coli bacteria alone (black lines) and E.coli bacteria incubated with RFPC + (dark grey lines) and RFPC -(light grey lines) at 2x10 -10 M in LB medium (A) or M9 medium (B). The error bars indicate the difference between the three replicates in the experiment (<6.8%).

Figure 4 . 9 .

 49 Figure 4.24 Red auto-fluorescence images of E.coli alone (100x objective, A: bright filed image, IT=500ms; B: fluorescence image, IT=1s). Additional images are shown in Figure S4.7 in Section 4.9. After incubation with RFPC + , instead of entering inside the bacteria and labeling the DNA, RFPC + labelled the bacteria membrane in a very homogeneous way (Figure 4.25).

Figure 4 .

 4 Figure 4.25 RFPC + labelling the membrane of E.coli bacteria (100X objective, A: bright field image, IT=500ms; B: fluorescence image, IT=1s). Additional images are shown in Figure S4.8 in Section 4.9. A similar observation happened to RFPC -, instead of labelling the cytoplasm of bacteria, RFPC -labelled the membrane of E.coli (Figure 4.26). Bothe RFPC -and RFPC + seems label the membrane and didn't enter inside bacteria. The reason for RFPCs labelling the bacterial surface membrane instead of entering inside the bacteria is the hydrophobic head of the RFPCs would insert and remain in the hydrophobic part of the phospholipid bilayer (Figure 4.27). Although the membrane is negatively charged, which in theory means that RFPC + should label the membrane better, it is difficult to tell the difference between the RFPC + and RFPC -in their ability to stain the bacterial membrane in these experimental condition. Both of them label bacterial membrane clearly and homogeneously.

Figure 4 .

 4 Figure 4.26 RFPC -labelling the membrane of E.coli bacteria (100X objective, A: bright field image, IT=500ms; B: fluorescence image, IT=1s). Additional images are shown in Figure S4.9 in Section 4.9.

Figure 4 . 27

 427 Figure 4.27 One hypothesis of mechanism for RFPCs labelling bacterial membrane.

  The bacterial cytoplasm was targeted by GFPC -(Figure4.28B) which is the green part, and the bacterial membrane was labelled by RFPC + (Figure4.28C) which is the red part. By overlapping the two images together (Figure4.28D), it appeared that the green part and the red part do not overlap except for the membrane boundary. This shows that GFPC -and RFPC + are compatible with each other and that they could be used for bacteria internal and external bioimaging at the same time.

Figure 4 .

 4 Figure 4.28 E.coli interacting with GFPC -and RFPC + (A: bright field image, IT=500ms; B: green fluorescence image, IT=5s; C: red fluorescence image, IT=1s; D: overlapped of image B and C).

Figure 4 . 19 )

 419 indicating to a non-uniform labelling. RFPCs are very easy and cheap to synthesize and are non-toxic to bacteria. Moreover, they can label the bacterial membrane by simple incubation in common growth medium (M9) which is very easy to handle compared to PKH26 which needs a special solvent for dilution. The spectra of PKH26 and RFPCs are displayed in Figure 4.29. Their maximum absorption and emission wavelength are shown in Table4.5. From the spectra, one can notice that PKH26 and RFPCs share very similar maximum absorption wavelength (PKH26: 551nm, RFPC + : 552nm, RFPC -: 553nm), however the maximum is located at different wavelength while the Stoke Shift of RFPCs (RFPC + : 1477 cm -1 , RFPC -: 1076 cm -1 ) are higher compared with the one of PK26 (818 cm -1 ). A larger Stoke shift between absorption and emission bands allows more efficient collection of emitted light[START_REF] Demchenko | Introduction to Fluorescence Sensing[END_REF] , which is better for imaging experiments.

Figure 4 .

 4 Figure 4.29 PKH26 (black lines) in PBS and RFPC + (dark grey lines) and RFPC -(light grey lines) in water (pH=7.0): absorption (full lines), fluorescence emission (dotted lines) normalized spectra.

  than PKH26. In order to test the minimum staining concentration for PKH26, I ran an experiment to decrease the quantity of PKH26 continually and to measure the extent of membrane staining by microscopy (Figure 4.30).

Figure 4 . 30

 430 Figure 4.30 Images of different concentration PKH26 labeling the membrane of E.coli (PKH26 concentration A: 2x10 -8 M, B: 2x10 -9 M, C: 2x10 -10 M).

Figure 4 .

 4 30A, the concentration of PKH26 is 2x 10 -8 M which is 100 times less than the concentration we used in previous experiments, and the staining is successfully made. However, when the concentration is decreased to 2x10 -9 M (Figure 4.30B), 2x10 -10 M (Figure 4.30C), the labeling is clearly less efficient. It is noticeable that we use the concentration of 2x10 -10 M for RFPCs as the staining concentration. In conclusion, we can use at least 100 times less RFPCs material to label the same amount of bacteria than a commercial membrane dye PKH26. And RFPCs are also very easy to handle during experiments without need of extra equipment or special conditions.

Figure 4 . 31

 431 Figure 4.31 Summary of FPCs and their bioimaging application of E.coli bacteria.

Figure S4. 1 Figure

 1 Figure S4.1 Fluorescence image (60X objective, IT=10s) of E.coli alone (A) and E.coli interacting with GFPC 0 (B) and GFPC -(C).A1B1

Figure S4. 3

 3 Figure S4.3 Images of DRAQ5 labeling the DNA of E.coli (60X objective, A: phase contrast image, IT=100ms; B: green fluorescence image, IT= 20s; C: red fluorescence image, IT=20s; D: overlapped image of A and C).

Figure S4. 4

 4 Figure S4.4 Bioimaging of E.coli bacteria incubated with GFPC -and DRAQ5 (60X objective A: phase contrast image, IT=100ms; B: green fluorescence image, IT= 20s; C: red fluorescence image, IT=20s). Additional images are shown in Figure S4.4 in Section 4.9.

Figure S4. 5

 5 Figure S4.5 Fluorescence images of PKH26 labeling the membrane of E.coli (100X objective, fluorescence image, IT=5s).

Figure S4. 6 E

 6 Figure S4.6 E.coli interacting with GFPC -and PKH26 (A: green fluorescence image, IT=5s; B: orange fluorescence image, IT=5s; C: overlapped image of A+B).

Figure S4. 7

 7 Figure S4.7 Red auto-fluorescence images of E.coli alone (100x objective, A: bright filed image, IT=500ms; B: fluorescence image, IT=1s).

Figure S4. 8

 8 Figure S4.8 RFPC + labelling the membrane of E.coli bacteria (100X objective, fluorescence image, IT=1s).

Figure S4. 9

 9 Figure S4.9 RFPC -labelling the membrane of E.coli bacteria (100X objective, fluorescence image, IT=1s).

  ] Hydrodynamic diameter (D h ) determined by DLS.

λ

  abs /nm λ em /nm Φ Hydrodynamic diameter (D h ) determined by DLS. [b] Error of 15% 182 . [c] Styrene conversion determined by gravimetry and corrected with the loss of styrene by evaporation 203 .

Figure 5 . 1 , 2 .

 512 Figure 5.1, their main spectroscopic data are given in Table 5.2. The particle suspensions have a maximum absorption at 488 and 490 nm and a maximum fluorescence emission at 519 and 521 nm in FANP1 and FANP2 respectively, in water at pH 8. There is a slight red shift compared to the free FA dye 201 which may come from the change of the local environment of the dye. The fluorescence quantum yield of the nanoparticles is between 12% and 14% for both particles at pH 8. This is lower than the free fluorescein amide dye (pH > 7, Φ F = 79% 177 ) which may be due to the formation of FA aggregates in the shell. The number of grafted fluorescein molecules is 530 and 340 for FANP1 and FANP2 respectively as estimated from the absorption spectra 204 .

Figure 5 . 1

 51 Figure 5.1 Normalized absorption (full lines) and fluorescence emission (dotted lines) spectra (λexc=495nm) of a suspension of FANP1 (grey lines) and FANP2 (black lines) in water at pH=8. The spectroscopic behavior of these FA-grafted nanoparticles was then studied at various pH values (Figure 5.2). The fluorescence spectra at pH=8 for FANP1 (Figure 5.2A) and FANP2 (Figure 5.2C) have a main band centred at 519 nm and 521 nm respectively. This main band was attributed to fluoresceinamide. The fluorescence spectra were then recorded at different pH (phosphate/citrate buffers 1mM in 140mM NaCl). Upon a decrease in the pH, the spectral intensity decreased both in the absorbance and fluorescence for FANPs. This behaviour is similar to that of fluorescein in water. Moreover the pKa was determined from the fluorescence spectra using the Henderson-Hasselbach equation:

Figure 5 . 2

 52 Figure 5.2 Fluorescence emission spectra of FANP1 (A) and FANP2(C) as a function of pH (λ exc =495nm; phosphate/citrate buffer 1mM and 140mM NaCl). B: Change in fluorescence intensity of FANP1 at 520nm as a function of pH. The data was fit using the Henderson-Hasselbach equation with pKa=6.49. pH values: 7.37, 7.07, 6.94, 6.41, 5.84, 4.95, 4.41, 3.85, 3.01. D: Change in fluorescence intensity of FANP2 at 520nm as a function of pH. The data was fit using the Henderson-Hasselbach equation with pKa=6.53. pH values: 7.80, 7.14, 7.01, 6.55, 6.27, 5.85, 5.07, 4.57, 4.00.

Figure 5 . 3

 53 Figure 5.3 Evolution of fluorescence emission spectra of FANP1 or FANP2 over four hours ( exc =495nm; Modified M9 Minimal Medium,). A and C are at room temperature and B and D are at 37°C. A spectrum was taken every 30mins.

Every 6 Figure 5 . 6

 656 Figure 5.6 Growth curves of E.coli bacteria alone and of E.coli bacteria incubated with FANP2 particles at 8.2x10 -7 M, 4.1x10 -7 M and 8.2x10 -8 M. The error bars indicate the difference between the three replicates in the experiment.

  bacterial growth increased the fluorescence intensity of both FANPs decreased significantly (Figure 5.7A, B). The fluorescence intensity of FANP2 for example decreased more than 80% during bacterial growth. Thanks to the calibration curve (Figure 5.2D) the pH can be evaluated by recording the change in fluorescence intensity of FANP2 in the growth medium (Figure 5.7C). It was found that these pH values were in full agreement with those measured by a pH meter (Figure 5.7E). Incubating FANP1 with E.coli gave similar results as those obtained with FANP2 (Figure 5.7D, F). In conclusion, the pH-sensitive nanoparticles can be used to precisely monitor the pH variation due to bacterial growth (Figure 5.7).

Figure 5 . 7

 57 Figure 5.7 Change in the fluorescence emission spectra of FANP2 (A) and FANP1 (B) with bacterial growth (λ exc =495nm; Modified M9 Minimal Medium). B: Change in fluorescence intensity of FANP2 (C) and FANP1 (D) at 520nm and corresponding pH of the growth medium calculated by the Henderson-Hasselbach equation. Real-time bacterial growth curve (dotted line) and the corresponding pH evolution as measured by a pH meter (grey line) or the corresponding pH change calculated by the Henderson-Hasselbach equation from the change in fluorescence (black crosses)for FANP2 (E) and FANP1 (F).

  37°C in the dark. Every 30 minutes the growth of the cells was monitored by using a spectrophotometer (Figure 5.8). At the same time, the emission spectra of the samples were measured. The corresponding decrease in the percentage of the fluorescence intensity of FA molecules, FANP1 and FANP2 as a function of time is shown in Figure 5.8.

Figure 5 . 8 Figure 5 . 9

 5859 Figure 5.8 Measurements of E.coli bacteria growth incubated with FA molecules (black lines), FANP1 (light grey lines) and FANP2 (dark grey lines) using either the measurement of OD@600nm (dotted lines-scale on the right axis) or the decrease (%) of fluorescence intensity (full lines-scale on the left axis) as a function of time for FA molecules, FANP1 and FANP2. The two lines represent the replicates within the experiment.

Figure 5 . 8 ,

 58 Figure 5.8, the fluorescence intensity measurements from the FANP1 and FANP2 particles are very reliable and reproducible from the very beginning until the end of the experiments.

5 . 11 .Figure 5 . 10 Figure 5 . 11

 511510511 Figure 5.10 Fluorescence evolution screening of FANP1 (A) and FANP2 (C) incubated with E.coli. The decrease (%) of fluorescence intensity as a function of time for three different concentrations of FANP1 (B) and FANP2 (D): 8.2x10 -7 M (black lines), 4.1x10 -7 M (dark grey lines) and 8.2x10 -8 M (light grey lines). Data were extracted from E.coli background fluorescence; the error bars indicate the difference between the three replicates in the experiment.

12 .

 12 high throughput experiments such as the detection of antibiotic resistance. In order to compare the results obtained by the change in OD with the change in fluorescence of FANPs(8.2x10 -8 M) the effects of several common antibacterial drugs (Chloramphenicol, Kanamycin, and Ampicillin) on bacteria growth and metabolism were measured. The concentration of antibiotics used were 0.5, 1, 5, 10, 50 µg/mL, which cover the range of the known recommended concentrations of these antibiotics. The inhibitory effect of these drugs on the growth of E.coli as measured by the increase of the OD or the decrease in fluorescence of FANP1 is shown in Figure5.When increasing the concentration of antibiotics, the growth rate of E.coli bacteria decreases as measured by OD (Figure5.12A, C, E). At the same time, the fluorescence intensity of FANP1 decreases when increasing the dose of the drugs (Figure5.12B, D, F).

Figure 5 . 12 Figure 5 . 13

 512513 Figure 5.12 The inhibitory effect of three antibiotics on the growth of bacteria: Chloramphenicol (A,B), Kanamycin (C,D), and Ampicillin (E,F) at different concentrations (0.5, 1, 5, 10, 50 µg/mL). The change in the growth of E.coli was measured by an increase of the OD (A, C, E) and by a decrease of fluorescence of FANP1 (B, D, F). The error bars indicate the difference between the three replicates in the experiment.

FANP1

  and FANP2 were added to the growth medium with an initial concentration of E.coli of 3-6x10 5 CFU/mL (OD=0.003-0.006) inside a cuvette (Figure5.14A a0, c0). After an overnight incubation, the fluorescence intensity of the cuvettes with bacteria decreased significantly (Figure5.14A a1, c1) and the decrease could be easily seen by eye with the help of a lamp (exc@470nm). However, the fluorescence intensity of the cuvettes without bacteria remains unchanged (Figure5.14A b0, b1, d0, d1).

Figure 5 .

 5 Figure5.14 A: a0, c0: FANP1 and FANP2 in growth medium with an initial concentration of E.coli of 3x10 5 CFU/ml (OD=0.003). b0 and d0: FANP1 and FANP2 in growth medium. a1, b1, c1, d1 are fluorescence images of a0, b0, c0, d0 after one night incubation (37°C). B: Fluorescence images of bacterial growth on FANP2 agarose (a0, b0, c0), compared to fluorescence images of FANP2 agarose alone (d0). a1, b1, c1, d1 are fluorescence images of a0, b0, c0, d0 after one night incubation (37°C). C: Fluorescence images of bacterial growth on FANP1 agarose (a0, b0, c0), fluorescence images of FANP1 agarose alone (d0). a1, b1, c1, d1 are fluorescence images of a0, b0, c0, d0 after one night incubation (37°C).

Figure 6 . 1

 61 Figure 6.1 Fluorescent nanomaterials for internal and external bioimaging and biosensing application on E.coli bacteria. (A) FNPs: Fluorescent nanoparticles for internal bioimaging of E.coli. (B) FNP-

  Red fluorescent polymer chains (RFPCs) could label the bacterial membrane easily by standard incubation (Figure6.1D). GFPC -and RFPC + were compatible with each other and they could be used for bacteria internal and external bioimaging at the same time.Compared with the commercial membrane dye PKH26, RFPCs have shown significant advantages: we can use at least 100 times less RFPCs material to label the same amount of bacteria.The designed FPCs series are highly water-soluble, biocompatible, stable and bright.With different multi-color image tests, FPCs have shown good compatibility with different types of commercial dyes. Moreover, they are very easy and cheap to synthesize and can be easily used to either internal (cytoplasm) or external (membrane) bioimaging applications of bacteria.After these bioimaging studies, two highly water-soluble, biocompatible, stable and bright fluoresceinamine based pH-sensitive nanoparticles (FANPs) were investigated for biosensing application on E.coli bacteria (Figure6.1E). They could rapidly and accurately detect bacterial growth by signalling the change of pH resulting from cellular metabolism.

1

 1 Synthesis of fluorescent nanoparticles (FNPs)BODIPY derivative: BODIPY methacrylate-1 (BDPMA1), and PEO-b-PAA-TTC-C 12 macroRAFT agents were synthesized as described elsewhere[START_REF] Grazon | Élaboration de nanoparticules fluorescentes à base de BODIPY par polymérisation RAFT en miniémulsion : synthèse, caractérisation et fonctionnalisation de surface[END_REF] . Reversible additionfragmentation chain transfer (RAFT) polymerization 215 of styrene and BDPMA1 is performed in the presence of poly(ethylene oxide)-b-poly(acrylic acid), PEO 45 -b-PAA 10 -TTC-C 12 (PEO:

Scheme 7 . 1

 71 Scheme 7.1 Synthetic pathway for fluorescent nanoparticles (FNP -). We obtained fluorescent nanoparticles PEO 45 -b-PAA-b-P(S-co-BODIPY)-TTC-C 12 with about 1750 chains, and 1.7 BODIPY per chain. The molar mass of the FNP -(negatively charged fluorescent nanoparticle) is 3x10 7 g/mol 27 .

Scheme 7 . 2

 72 Scheme 7.2 Synthetic pathway to get positively charged FNP (R refers to Ethylenediamine or Poly(ethylene glycol)diamine).

Scheme 7 . 3

 73 Scheme 7.3 Synthetic pathway for biotin conjugation to FNP -.

Scheme 7 . 8

 78 Scheme 7.8 Synthetic pathway for positive red fluorescent polymer chain (RFPC -)

  Scheme 7.10 Preparation of pH nanosensors (FANP1 and FANP2) from PSNP1 and PSNP2.

1

 1 Absorption and fluorescence spectroscopyUV-visible spectra were measured on a Varian Cary 5000 (Palo Alto, CA USA) double beam spectrometer using 4.5mL BRAND Polymethacrylate (PMMA) Cuvette. Excitation and emission spectra were recorded on a SPEX SPEX Fluoromax-3 (Horiba Jobin-Yvon). A rightangle configuration was used. Optical density of the samples was checked to be less than 0.1 to avoid reabsorption artifacts. The relative fluorescence quantum yields Φ F were determined by using Rhodamine 590 (Φ F = 0.95 in ethanol) or Rhodamine 101 (Φ F = 0.96 in ethanol) as a reference (error of 15%)[START_REF] Kubin | Fluorescence quantum yields of some rhodamine dyes[END_REF]180 .

7. 3 . 8

 38 Photostability and reversibility assessment of FANPs 2.5mL of Modified M9 minimal growth medium were placed inside a 4.5mL BRAND Methacrylate (PMMA) Cuvette. Aliquots of either FANP1 or FANP2 solutions were added to reach a final concentration of 8.2x10 -7 M. Samples were incubated at either 37°C or at room temperature in the dark and the fluorescence emission spectrum (λ exc =495nm) of each sample was monitored using a SPEX Fluoromax-3 (Horiba Jobin-Yvon) every 30 minutes. The experiments lasted for 4 hours. The reversibility and dynamic behaviour of FANPs was investigated by alternating the pH of the modified M9 minimal growth medium between pH 4 and 8 several times inside the cuvette in the presence of FANPs by adding 1M NaOH or HCl alternatively. An overnight experiment was also carried out to assess the photostability of FANPs. 150µL of the Modified M9 minimal growth medium were placed in each well of a 96 well Falcon Polystyrene Flat Bottom Plate. Different volumes (from 2 to 10 µL) of the stock FANP1 and FANP2 solutions were added into the solution to reach final FA concentrations of 8.2x10 - 7 M, 4.1x10 -7 M and 8.2x10 -8 M. Each plate contained three repeats of the same concentration.One control with only the Modified M9 minimal growth medium was also prepared. 70 µL of mineral oil were added to each well in order to avoid evaporation. Samples were incubated in a plate reader (Perkin Elmer Victor3 1420 Multilabel Plate Counter) at 37°C in the dark.

7 . 4 . 2

 742 For pH-sensor experiments described in Chapter 5, I use a Modified M9 minimal medium with a lower concentration of phosphate salts in order to decrease the buffering capacity of the growth medium and thus obtain a more sensitive measure of bacterial growth from the response of the FANPs. The Modified M9 minimal medium contains 5.9mM Na 2 HPO 4 •2H 2 O, 4.4mM KH 2 PO 4 , 3.7mM NH 4 Cl, 1.7mM NaCl, 2mM MgSO 4 , 0.1mM CaCl 2 , 19.6µM tryptophan, 20.6µM thymidine, 0.5% casamino acids, 22.2mM glucose. Toxicity assessments of FNPs and FPCs on E.coli Live E.coli bacterial cells (K-12 BW25113) were used for the following experiments. Bacterial cultures were prepared overnight from stock cultures inoculated in Luria Broth (LB) growth medium.The overnight culture of bacteria was diluted 1:1000 in two different growth media: M9 or LB in order to check the growth medium influence. 150µL of bacterial solution were placed in each well of a 96 well Falcon Polystyrene Flat Bottom Plate. Different volumes of the FNPs or GFPCs or RFPCs solutions were added into the E.coli solution to reach final concentration of 6.6x10 -3 M (FNPs), 6.6x10 -5 M (GFPCs) or 2x10 -10 M (RFPCs). Each plate contained three repeats of the same concentration. One control with only bacteria and one blank with only M9 minimal growth medium or LB medium were also prepared. 70 µL of mineral oil were added to each well in order to avoid evaporation. Samples were incubated in a plate reader (Perkin Elmer Victor3 1420 Multilabel Plate Counter) at 37°C in the dark.The growth of the cells was monitored every nine minutes by reading the optical density (OD@600nm). The experiments lasted overnight.

1

 1 Chemical treatment to introduce FNPs in E.coli Bacteria 1mL of LB growth medium with an initial concentration of 5x10 7 CFU/mL (OD=0.5) of E.coli bacteria was placed inside a 2mL Eppendorf Safe-Lock Tube. Different volumes of FNP - solutions were added to this E.coli suspension to reach different final concentrations:

  Tween 20 is added to promote more effective washings, resulting in decreased non-specific background staining) once and suspended in 20 µL before bioimaging experiments.

7. 7

 7 Incubation to introduce FPCs into E.coli bacteria 7.7.1 Incubation to introduce GFPCs into E.coli bacteria

7. 7 . 2

 72 Incubation to introduce RFPCs into E.coli bacteria1mL of M9 growth medium with an initial concentration of 5x10 7 CFU/mL (OD=0.5) of E.coli bacteria was placed inside a 2mL Eppendorf Safe-Lock Tube. Different volumes of RFPC + and RFPC -solutions were added to this E.coli suspension to reach a final concentration of 2x10 -10 M. Two repeats of the same concentration were carried out during the experiments.

7. 8 Figure 7 . 1

 871 Figure 7.1 Flow Cytometry (BD, FACS Calibur) (A), analysis of flow cytometry (B) X axis: green fluorescence, Y axis: red fluorescence. R1 refers to live non-fluorescent cells percentage, R2 refers to live green fluorescent cells percentage, R3 refers to dead red-green fluorescent cells percentage and R4 refers to dead red fluorescent cells percentage.Live cells have intact membranes and are impermeable to dyes such as propidium iodide (PI), which only leaks into dead cells with compromised membranes. In this way, we could measure the percentage of live cells in each sample, and assess the toxicity of nanoobjects on E.coli 217 . Before flow cytometry, propidium iodide (PI, 10µL, 1.3x10 -2 mg/mL) was added into each sample. PI emits in the red (317nm) once it is bound to the DNA inside the bacteria. FNPs or GFPCs emit in the green (548nm), the emission signal will be measured if they remain inside or attached to the membrane of the bacteria, otherwise they are lost during the washing steps. Analysis of the cells by flow cytometry is shown in Figure 7.1. R1 refers to live non-fluorescent cells percentage, R2 refers to live green fluorescent cells percentage, R3 refers to dead red-green fluorescent cells percentage and R4 refers to dead red fluorescent cells percentage.

7. 9 7 . 9 . 1 7 . 9 . 2

 9791792 Techniques for rapid detection of Escherichia coli growth by fluorescent pH-sensitive nanoparticles (FANPs) Screening of bacterial growth in a 96 well plate Live E.coli bacterial cells (K-12, BW25113) were used for the following experiments. Bacterial cultures were prepared overnight from stock cultures inoculated in Luria Broth (LB) growth medium. The overnight culture of bacteria was diluted 1:1000 in the Modified M9 minimal growth medium. 150µL of this bacterial solution were placed in each well of a 96 well Falcon Polystyrene Flat Bottom Plate. Different volumes (from 2 to 10 µL) of the stock FANP1 and FANP2 solutions were added into the E.coli solution to reach final concentrations of 8.2x10 - 7 M, 4.1x10 -7 M and 8.2x10 -8 M. Each plate contained three repeats of the same concentration. One control with only bacteria and one blank with only the Modified M9 minimal growth medium were also prepared. 70 µL of mineral oil were added to each well in order to avoid evaporation. Samples were incubated in a plate reader (Perkin Elmer Victor3 1420 Multilabel Plate Counter) at 37°C in the dark. The growth of the cells was monitored every 6 minutes by reading the optical density (OD@600nm). Fluorescence was measured using a F485/14 filter for excitation and a F535/40 filter for emission. The experiments lasted overnight for ten hours. Real-Time detection of E.coli 2.5mL of Modified M9 minimal growth medium with an initial concentration of 2x10 7 CFU/mL (OD=0.2) or 3-6x10 5 CFU/mL (OD=0.003-0.006) of E.coli bacteria were placed inside a 4.5mL BRAND Methacrylate (PMMA) Cuvette. Different volumes of FA (fluoresceinamine) solution, or FANP1 or FANP2 suspensions were added to this E.coli suspension to reach a final concentration of 8.2x10 -7 M. Two repeats of the same concentration were carried out during each of the experiments.

FANP2 7 . 9 . 5

 795 solutions were added to reach a final concentration of 8.2x10 -7 M. Stirring for 2 minutes yielded a homogeneous mixture which was poured into each of the 1cm wells of a Nunc 4 well cell culture plate to form am agarose layer upon cooling to room temperature. E.coli bacterial solutions with an initial concentration of 1-2x10 4 CFU/mL were added on different positions of the culture plates. The samples were incubated at 37°C in the dark overnight. Imaging of E.coli growth Fluorescence images of the samples in the cuvettes and in the culture plates were taken by a smartphone (Google Nexus 5) by putting samples on a Safe Imager™ 2.0 Blue Light Transilluminator (exc@470nm).

Figure 7 . 2 7 . 10 . 5

 727105 Figure 7.2 Nikon inverted microscope ECLIPSE TI-E (A), Scheme of Nikon epifluorescence microscope (B) The scheme of the Nikon Epifluorescence microscope (C-HGFI Intense light Nikon) is shown in Figure 7.2B. The resolution of the microscope is about 450 nm at the wavelength of 528 nm. FITC-3540C BrightLine single-band filter (Figure 7.3A) is used in this microscope. It provides both high brightness and signal-to-noise ratio. It is also ideal for fluorophores that have narrow, more closely spaced excitation and emission peaks, like BODIPY. E.coli cells with FNPs or GFPCs are illuminated with light of a wavelength (482 nm) which excites FNPs or GFPCs. TRITC-B BrightLine single-band filter (Figure 7.3B) is also used for RFPCs. E.coli cells with RFPCs are illuminated with light of a wavelength (543 nm) which excites RFPCs. The fluorescent light is then imaged through a microscope objective with an emCCD camera. Objectives 60X (CFI S Plan Fluor ELWD, NA 0.7) and 100X (CFI Plan Apo VC, NA 1.4, Nikon 50 TYPE NF IMMERISION OIL) are both used in this research. During the experiment, I take intensity images of the samples. Firstly, the focus is made on the bacteria with white light and the phase contrast image is recorded. Secondly, I turn on the mercury lamp and choose the right filter for the fluorescence image. We can also overlap the phase contrast and

BODIPY methacrylate- 1 (

 1 photophysical properties of BODIPY chromophores are known to be weakly affected by their environment. Hence BDPMA1 conversion was determined as:

  f a : The fraction of the initial fluorescence of FNP -or GFPC -that is accessible to quencher methylene blue. K a : The Stern-Volmer quenching constant of the accessible fraction.[Q] : the concentration of quencher methylene blue. This modified form of the Stern-Volmer equation allows f a and K a to be determined graphically. A plot of F 0 /∆F versus 1/[Q] yields f a -1 as the intercept and (f a K a ) -1 as the slope. A y-intercept of f a -1 may be understood intuitively. The intercept represents the extrapolation to infinite quencher concentration (1/[Q] = 0). The value of F 0 /(F 0 -F) at this quencher concentration represents the reciprocal of the fluorescence that was quenched. At high quencher concentration only the inaccessible residues will be fluorescent. Brightness 27 Brightness is the total efficiency of absorption and fluorescence emission of a fluorophore. Brightness of a dye depends on two properties: molar extinction coefficient at the excitation wavelength ε(λ) and fluorescence quantum efficiency, Φ F 26 . The brightness (B) of a fluorophore is proportional to the molar extinction coefficient at the excitation wavelength ε(λ) and quantum yield (Φ F ), as indicated in the following relationship 26 : 𝐵 = 𝜀(𝜆) × 𝛷 𝐹 For fluorescent nanomaterials (FNPs or GFPCs), the brightness is defined as 27 : of fluorophores contained inside the FNPs or GFPCs ε(λ): molar extinction coefficient of the fluorophore at the excitation wavelength, we assumed that ε 𝑚𝑜𝑛𝑜𝑚𝑒𝑟 528 ≈ ε 𝑝𝑜𝑙𝑦𝑚𝑒𝑟 528 (73000 L.mol -1 .cm -1 ) 27 . This assumption appears acceptable since the photophysical properties of BODIPY chromophores are known to be weakly affected by their environment. Φ F : fluorescence quantum yield of the FNPs or GFPCs. 76. Llopis, J., McCaffery, J. M., Miyawaki, A., Farquhar, M. G. & Tsien, R. Y. Measurement of cytosolic, mitochondrial, and Golgi pH in single living cells with green fluorescent proteins. Proc. Natl. Acad. Sci. 95, 6803-6808 (1998). 77. Westermann, B. & Neupert, W. Mitochondria-targeted green fluorescent proteins: convenient tools for the study of organelle biogenesis in Saccharomyces cerevisiae.

Table 2 . 1

 21 Main spectroscopic properties of BDPMA1 and FNPs

	Sample	Structure	λ abs /nm	λ em /nm	Φ F /% water [a]

Table 3 .

 3

			Sample	λ abs	λ em	Φ F /%	Brightness/	D t /nm [c]	D h /nm [d]
						/nm	/nm	PBS [a]	M	-1 cm -1[b]
			FNP	-	529	543	36	7.8x10	7	65	65
			FNP	--biotin	529	543	35	7.6x10	7	67	67
	FNP	--biotin-streptavidin	529	543	34	7.4x10	7	77	78
	FNP	--biotin-streptavidin-	529	543	30	6.5x10	7	97	92
			antibody
	[a] Error of 15%

1 Main spectroscopic properties of FNP -, FNP --Biotin, FNP --Biotin-streptavidin, FNP --Biotinstreptavidin-antibody 182 ; [b] Brightness is determined by Equation 7.5 in Chapter 7; [c] theoretical diameter (D t ); [d] hydrodynamic diameter (D h ) determined by DLS.

Table 3 . 2

 32 Main spectroscopic properties of FNPB, FNPB-antibody

	Sample	λ abs	λ em	Φ F /%	Brightness/
		/nm	/nm	PBS [a]	M	-1 cm -1[b]
	FNPB	529	541	42	5.3x10	7
	FNPB-antibody	529	541	28	3.5x10	7
	[a] Error of 15%

182 ; [b] Brightness is determined by Equation 7.5 in Chapter 7.

Table 4 .

 4

		1 Kinetic study of the synthesis of GFPC -and GFPC°
	Sample	Time/mins BDPMA1
		conv./% [a]

Table 4 .2 Critical micelle concentration(CMC) for GFPCs

 4

	CMC/ mg.ml -1	20°C/ water	37°C/ LB
	GFPC -	0.3	0.4
	GFPC 0	0.38	0.5
	A	B
	C	D

Table 4 . 3

 43 Main spectroscopic properties of BDPMA1 and GFPCs

				water			toluene
	Sample	Structure	λ abs	λ em	Φ F	λ abs	λ em	Φ F	BDPMA1	Brightness	D t [c]	D h [d]
			/nm	/nm	/% [a]	/nm	/nm	/% [a]	per chain	/M	-1 cm -1[b]	/nm	/nm
	BDPMA1	-	-	-	-	528 540	70	-	51100	-	-
	GFPC°	P(APEG 17 -co-	528 548	29	528 542	78	3	63510	4-6	142
		BDPMA1 3 ) 20 -TTC-C 12
	GFPC-	P(AA 9 -co-APEG 9 -co-	528 548	20	528 542	74	2	29200	4-6	84
		BDPMA1 2 ) 20 -TTC-C 12

[a] Error of 15%

Table 4 . 4

 44 Main spectroscopic properties of BDPMA2 and RFPCs

				water			ethanol
	Sample	Structure	λ abs	λ em	Φ F	λ abs	λ em	Φ F	D t [b]	D h [c]
			/nm	/nm	/% [a]	/nm	/nm	/% [a]	/nm	/nm
	BDPMA2	-	-	-	-	552 594	53	-
	RFPC +	P(DMAEA-APEG-co-	552 601	12	550 585	32	4-6	30
		BDPMA2)-TTC-C 12
	RFPC -	P(AA-co-APEG-co-	553 588	10	551 587	31	4-6	17
		BDPMA2)-TTC-C 12
	[a] Error of 15%

182 , [b] theoretical diameter(D t ) [c] hydrodynamic diameter (D h ) determined by DLS in water.

Table 4 . 5

 45 Maximum absorption and emission wavelength of PKH26 and RFPCs

		λ abs /nm	λ em /nm	Stoke Shift/ cm -1
	PKH26	551	577	818
	RFPC +	552	601	1477
	RFPC -	553	588	1076

Table 5 . 1

 51 Characterization of nanoparticles (PSNPs) having different shells[START_REF] Grazon | Élaboration de nanoparticules fluorescentes à base de BODIPY par polymérisation RAFT en miniémulsion : synthèse, caractérisation et fonctionnalisation de surface[END_REF]

	Sample	Shell	Conv.s	[a]

Table 5 . 2

 52 Spectroscopic properties of FANPs in water (pH=8)[START_REF] Grazon | Élaboration de nanoparticules fluorescentes à base de BODIPY par polymérisation RAFT en miniémulsion : synthèse, caractérisation et fonctionnalisation de surface[END_REF]

	Sample	Shell	D h /nm	[a]

  Husson, Christian Jean-Baptiste, Jacky Fromont, Arnaud Brosseau, Stéphanne Maisonnneuve, Laurent Galmiche, Ni Ha Nguyen, Laura Nodin, Meriem Stamboul, Claire Déo, Corentin Pavageau, Charlotte Rémy, Marine Louis, Paul Rouschmeyer, Clarisse Tourbillon, Jérémy Bell, Jérémy Malinge, Hervé Leh, Marie-Christine Bouger, Gladys Mbemba, Marine Baudin… I appreciate your warm help and kind assistance. I am deeply grateful for every colleague, you guys have always been so nice to me, and my work would not have been possible finished without the collaboration, help, and support of all of you. I also want to thank all my dear beloved friends I met in France during the past five years: Jia Su, Qing Zhou, Yayang Tian, Qing Zhang, Yanhua Yu, Yinbin Ruan, Yuanyuan Liao, Na Chen, Yuan Li, Sha Li, Lve Huang, Xiaoju Ni, Feifei Liang, Xiaoqian Xu, Tiantian He, Wei Song, Marta Gordel, Aleksandra Delplanque, Akiko Hatakeyama, Claudia Irene Trainito, Gaïa Grassini, and Rawand Masoud… You guys have lighten my life and made my stay in France so wonderful.

‡ LCP laboratory (UMR 7610), UPMC Univ Paris 06, France

Acknowledgements

My time at PPSM and LBPA has been impactful, formative, and an extraordinary experience that I will remember with gratitude throughout my life. I must express my great appreciation to these who accompanied me during my PhD study. My deepest gratitude goes first and foremost to my supervisors: Pr. Rachel Méallet-Renault, Dr. Bianca Sclavi, and Dr. Gilles Clavier for giving me the opportunity to do this research in ENS-Cachan. Words are simply not enough for expressing gratitude towards them. They are the one who are always available for research questions, and always ready to give advice and to help in overcoming difficulties either in the study or in life during my staying in France. They inspired me to do my best in performing experiments, preparing talks and writing report, also offer me many opportunities to participate in conferences and summer school. It is their intellectual support, encouragement, enthusiasm, and intelligent guidance, which made this dissertation possible. I want to express my warm and sincere thanks to Dr. Chloé Grazon, who helps me to get familiar with the lab, taught me hand by hand during experiments. I would not have completed my research without her encouragement and sound advice. I am very grateful to Jean-Frédéric Audibert, who was very experienced in microscope and he provides great assistant for my microscopy experiments, thanks to his useful discussion, patience and inspiration. I would like to express my great thanks to my dear be loved friend, Dr. Elisa Brambilla, who not only helped me a lot in biological experiments but also always warmhearted to help me out of any difficulties during my stay in Cachan. I would like to extend my gratitude to my committee: Pr. Niko Hildebrandt, Dr. Marie Erard and Dr. Romain Briandet for taking the time to read my dissertation and give me useful advice to improve my dissertation. I warmly thank to all the teachers during my master and PhD study, thank you for teaching me systemic science knowledge, you guide me on the way to science. I would like to acknowledge Dr. Fabien Miomandre, Dr. Malcolm Buckle, Dr. Eric Deprez, Dr. Isabelle Leray, Prof. Keitaro Nakatani, Prof. Pierre Audebert, Prof. Robert Pansu, Prof. Joanne Xie, Dr. Clemence Allain, Dr. Valerie Genot, Prof. Anne Debarre, Dr. Carine Julien, Dr. Guillaume Laurent, Dr. Remi Metivier, and Dr. Claude Nogues who helped me during my whole stay in Cachan. Thank you for your helpful suggestions and constant encouragement.

Synthesis of red fluorescent polymer chains (RFPCs)

The synthesis of BODIPY Methylnaphtalene-2 (BDPMA2) is described elsewhere [START_REF] Grazon | Élaboration de nanoparticules fluorescentes à base de BODIPY par polymérisation RAFT en miniémulsion : synthèse, caractérisation et fonctionnalisation de surface[END_REF] .

Negative red fluorescent polymer chain is synthesized in 1.4-dioxane at 80°C under nitrogen atmosphere. APEG (9 equiv.), AA (9 equiv.), TTCA RAFT agent (1 equiv.), DMF (5 equiv.), BDPMA2

Usage of PKH26 for labelling the membrane of E.coli bacteria

The following procedure uses a 500 µL final staining volume containing final concentration of 2x10 -6 M of PKH26 and 1x10 8 cells/mL. After introducing GFPC -into E.coli bacteria, samples were washed twice by PBS and re-suspended in 250 µL diluent C (an aqueous solution designed to maintain cell viability) in order to further perform membrane