FIGURE 1.5. Schematic representation of the helical structure of a self-assembled BTA-fiber. Figure reproduced from [7].
1.2.2 Applications of water-soluble BTA nanofibers
Considerable research has been performed about application areas for BTA nanofibers. Some future applications in for example regenerative medicine are mentioned in literature, where arti-ficial structures can be placed in the human body and be safely eliminated via biodegredation.[4]
Another application could be found in the field of electronics, in the form of conducting nanofibers.
More typical applications can be seen in figure 1.6, and involve nucleating agents for polymers, liquid crystals, organogels, MRI contrast agents, and microcapsules for drug delivery.[7]
FIGURE1.6. Examples of multiple different application areas for BTAs. Figure repro-duced from [7].
1.3. VISUALIZATION TECHNIQUES
1.3 Visualization techniques
For possible applications, more understanding of the properties of BTA-fibers is needed. It is necessary to visualize the polymers to study these properties. There are different visualization techniques available for imaging BTA-fibers.
One type of technique is cryo-TEM, a form of transmission electron microscopy, where samples are studied at cryogenic temperatures. When imaging BTA-fibers, the highly dynamic polymers are frozen during sample preparation, resulting in an image shown in figure 1.7. It is a very efficient method to look at the structure of BTA-fibers in high resolution. Long thin fibers in the order of micrometers in length and with a diameter of approximately 5 nanometers are observed.[6] However, due to the use of cryogenic conditions it is not possible to look at the fiber in its native state.
FIGURE1.7. Imaging BTA-fibers using cryo-TEM. Figure reproduced from [6].
An alternative method for imaging BTA-fibers, is using fluorescence microscopy. This is a common approach for studying dynamic events, compared to other methods that only provide a static snapshot view.[9] It is an accessible technique that does not require extreme imaging conditions. On the other hand, it makes use of a fluorescent dye molecule that needs to be coupled to the sample. Because of its high selectivity towards labeled fluorescent molecules, fluorescence imaging has become one of the most important imaging tools in biology.[10] An example of BTA-fibers imaged using fluorescence microscopy is shown in figure 1.8, where a cluster of fibers is visible.
FIGURE1.8. Imaging BTA-fibers using fluorescence microscopy.
The specificity of fluorescence microscopy is based on the difference between the wavelengths of absorbed light from a photon source and emitted light from the fluorophore. This is called the Stokes shift and it is schematically depicted in figure 1.9. The photon absorption process consists of a transition to an excited state of a fluorophore. After vibrational relaxation, the fluorophore returns to its ground state by fluorescent emission that can be measured. Using this principle, single fluorescent molecules are visible if the background has no autofluorescence.[10]
Total internal reflection fluorescent microscopy (TIRF), makes direct imaging of processes within very close proximity to the coverslip possible.[ 11] An evanescent field of excitation light, see figure 1.10, is created after excitation at a critical angle of the sample.[9] What follows is that the excitation depth is limited to approximately 100 nm, because of rapid decay of the excitation light intensity into the sample. This allows selective imaging of samples in close proximity to the surface. Combining the use of TIRF-illumination with state-of-the-art equipment, gives the ability to image single molecules.[12]
Fundamental limitations impede imaging at great resolution. The Abbe diffraction limit restricts the distinction between two objects separated by a distance less than approximately half the wavelength of light used to image the sample. This limit is roughly 250 nm, which is substantial compared to the size of BTA-fibers. In order to improve the resolution and to be able to study dynamic behaviour inside these fibers, we need to use an alternative visualization technique.
1.4. SUPER-RESOLUTION MICROSCOPY
FIGURE 1.9. Difference between the wavelength of absorbed and emitted light; the Stokes shift.
FIGURE1.10. Evanescence field created due to TIRF-angle.
1.4 Super-resolution microscopy
Recently, super-resolution microscopy techniques have been developed that are able to achieve a resolution up to 20 nm using visible light.[13] In these methods, the position of single fluo-rescent molecules is determined by fitting a two-dimensional Gaussian profile to the individual point-spread function of the excited molecule. Because this visualization technique is based on the detection of single molecules, it offers great improvement in resolution when compared to conventional microscopy.[14]
A particular technique called stochastic optical reconstruction microscopy (STORM), can
be used for high precision imaging. It makes use of sequential and stochastic reconstruction, resulting in precise localization of multiple single molecules.[14] The working principle of STORM entails the reconstruction of images after localization of a single photoswitchable dye. When only a fraction of the fluorophores are switched on, detection of a single fluorescent molecule is possible.[15] Using STORM, only a small, optically resolvable fraction of the photoswitchable dye is detected every snapshot. A photoswitchable dye is a fluorescent molecule that can be switched on and off. Cyanine dyes are an example of such fluorophores. They can be turned on and off reliably for hundreds of cycles before photobleaching.[15]
The switching between on and off state is a reversible photoconversion between a fluorescent and a dark state.[16] The exact mechanism of this photoconversion process is not known, but observations suggest that this switching is facilitated by a primary thiol.[17] For the commonly used Cy5 cyanine dye, the proposed model is the formation of a dark state, because of a thiol-addition to the polymethine bridge of the molecule.[16] This is a reversible chemical reaction, as can be seen in figure 1.11. Experiments showed that adding a primary thiol to the buffer with oxygen scavenging system improves on and off switching of the dyes.[18] Reducing the level of oxygen is important to prevent photobleaching.[9]
FIGURE1.11. Proposed model for the dark state by addition of primary thiol.
As explained before, during imaging, a Gaussian fit is performed for determining the exact place of emission after detection of a single fluorophore. Subsequently, many of such localizations can be combined to reconstruct a final image of high resolution.[15] Labeling BTA-monomers with a fluorophore like Cy5 thereby gives us the ability to look at BTA-polymer fibers. A comparison in achieved resolution between conventional and super-resolution techniques can be seen in figure 1.12, in which BTA-fibers imaged using conventional techniques look blurred compared to the super-resolution images.
In previous experiments, it was already found that monomers dynamically exchange position between different BTA-polymers, as shown schematically in figure 1.13, in contrast to covalently bonded polymers.[20] STORM was used to further examine this process. Experiments showed a homogeneous exchange of monomers along the polymer backbone.[19] By imaging the migration of Cy3-labeled and Cy5-labeled monomers between BTA-fibers, dynamic exchange between solvent and polymer-backbones was demonstrated. This investigation was performed using
1.5. SURFACE CHEMISTRY
FIGURE1.12. Difference in conventional imaging (top) and super-resolution imaging (down) of BTA-fibers. Figure reproduced from [19]
snapshot images at different time points after mixing, as can be seen in figure 1.14. Limitations in sample preparation prevent real-time imaging of these dynamics. The properties of the microscope glass coverslips favour fiber attachment and thus prevent us from imaging kinetic behaviour. Modification of these surfaces using surface chemistry could prove useful for looking at these dynamics, since anti-fouling coatings can be used to prevent settlement of organic residues.[21][22][23]
1.5 Surface chemistry
1.5.1 Basic concepts of silanization
Modification of the chemical composition of surfaces can be performed to obtain surface coatings with altered physical properties. This can be done by incorporating specifically selected functional molecules.
One type of organic molecules that can chemisorb to silicon oxide substrates (glass), are silanes. They contain a chemical group that can bind spontaneously and covalently to the surface and an organic rest group, schematically shown in figure 1.15. The silane part consists of a silicon atom with three leaving groups Xiand a linker part, which often consists of a long hydrophobic alkyl chain. Leaving groups are often hydroxy (−OH), chlorine (−Cl), methoxy (−OCH3) or ethoxy (−OCH2CH3).
Silanes can react with silanol groups ( −SiOH) on silicate substrates like glass surfaces.[ 25]
FIGURE1.13. Monomer exchange after mixing different BTA-fibers. Figure reproduced from [20]
This reaction is called silanization and is shown in figure 1.16. It contains a crucial condensation step, in which small molecules are released. This release prevents the reverse reaction. For example, when a trifunctional silane binds to the silanol groups on the surface, it releases three HX molecules per silane. When X is a hydroxyl group, three molecules of water are released.
The process of silanization consists of several underlying steps. To react, the molecules have to diffuse towards the substrate and at a certain point adsorb to the surface. After adsorption the molecules migrate laterally until they either desorb again or form a chemical bond with the substrate by binding in a potential minimum. The groups X of the silane will first react with water to form silanol groups (SiOH). This is called hydrolysis and it determines the overall reaction speed. The reaction rate depends on the identity of groups X and increases in the series OEt < OMe < Cl. After hydrolysis, the second step is a condensation reaction where the silanol group of the reactant binds to the silanol group on the surface, releasing a water molecule. The quality of deposition is strongly influenced by migration, possible nucleation to the substrate and desorption of the reactant.[24]
These processes are heavily influenced by the experimental steps of the functionalization protocol. Silane chemistry applied to silicate surfaces is usually performed using either vapor phase deposition or solution phase deposition of reactant silanes. Reactions take place at the gas-solid interface with deposition of thin films using chemical vapor deposition.[26] Especially at low pressure, surfaces can be coated with a homogeneous layer. A stable coating is obtained when a gas can react with the substrate surface, also forming gaseous by-products. Using this method, layer thicknesses of the order of 5 - 10µm can be obtained, depending on the exact system and
1.5. SURFACE CHEMISTRY
FIGURE1.14. STORM-imaging of fiber-exchange at different time points. Figure repro-duced from [19]
FIGURE1.15. Structure of a silane. Figure reproduced from [24].
protocol used.[24] The reaction can also be performed in solution, using solution phase deposition, in which we look at the liquid-solid interface. Surface coating can be obtained by immersing glass slides in a solution of reactant silanes. An incubation period of 10 - 60 minutes allows the dissolved silanes to react with the surface, resulting in a more densely functionalized surface.
1.5.2 Surface structure
The surfaces that are formed after reaction of silanes with solid substrates are often so-called monolayers. Because the binding of these molecules occurs spontaneously, these layers are called self-assembled monolayers (SAM).[27]
The growth of a self-assembled monolayer can take place in multiple ways and the quality of
FIGURE1.16. Reaction scheme for silanization. Figure reproduced from [24].
a SAM is highly dependent on the formation mechanism. Different growth mechanisms can be envisioned for these SAMs, because the formation of a SAM is governed by the aforementioned diffusion and adsorption processes. Those two processes occur at different rates, causing SAM formation to be either diffusion or adsorption controlled.[24] Different possible scenarios of monolayer growth are shown in figure 1.17. One is that reactants prefer to adsorb to aggregates already present on the substrate surface until a closed film is formed, scenario A. Another scenario is that first a closed film is formed, either ordered or disordered, scenarios B and C, after which the molecules in the film reorganize to form the final SAM in a second step.[24]
FIGURE1.17. Schematic view of growth of a monolayer by different scenarios. Figure reproduced from [24].
1.6. AIM AND OUTLINE OF THIS THESIS
The final structure of this monolayer is determined by three types of interaction.[24] The first one is the strength of the bond between the reactive head group and the substrate surface.
Further interactions are the attractive van der Waals forces between the linker groups of the silanes. The third and final interaction is the often repulsive interaction between the tail groups of the silanes due to, for example, steric hindrance.
A perfect monolayer is very difficult to form, because of the complex interplay between aforementioned interactions. It is found that it is sterically impossible to form a 1:1 stoichiometric saturation of all silanol groups on the substrate surface.[ 24] The hypothesis is that the surface layer will consist of a polysiloxane network anchored to the surface on some points, where approximately one in five silanes will bind to the surface and the other four are somewhat cross-linked above it, following scenario A, as can be seen schematically in figure 1.18.
FIGURE1.18. Schematic drawing of silanes bound to a silicon oxide surface. Figure reproduced from [24].
To prevent polymerization, water needs to be excluded from the reaction, hence the fact that usually this reaction is performed in dry organic solvents. However, a trace of water is needed for activating both the surface and the silanes. Water can react to the silanes in solution, forming polysiloxane networks that will precipitate, because the water molecules can compete with the hydroxyl groups on the substrate.[24] This could result in a rough surface, while the polysiloxanes can still, partly, bind to the substrate surface. Suitable silane molecules and reaction conditions should therefore be chosen to obtain the desired surface properties.
1.6 Aim and outline of this thesis
In this project, we aim to use surface chemistry approaches to create functionalized glass coverslips suitable for dynamic STORM experiments. The exchange of monomers between solution and aggregate state of a BTA-fiber in an aqueous environment can then be investigated.
In earlier experiments, BTA-fibers were physically adsorbed to glass microscope coverslips before imaging.[19][20] Once the fibers become physisorbed, no dynamic behaviour and monomer
exchange can be studied. This inhibits us from looking in real-time at changes in molecular composition of BTA-fibers. In order to understand how this exchange occurs, physisorption of BTA-fibers needs to be prevented. The approach towards imaging dynamic exchange of monomers consists of several experiments.
The first step is modification of the glass coverslip surface to prevent physisorption of BTA-fibers. The second step involves mixing in a bi-functional linker molecule, that at certain points links the BTA-fiber to the surface to prevent drift, while still maintaining its dynamic properties.
A schematic visualization of this approach is shown in figure 1.19. It shows that BTA-fibers are not completely bonded to the surface, but merely a loose flexible connection via linker molecules is formed, resulting in a fiber that still has the capacity to exchange monomers between solution and aggregate-state. The final step then consists of studying real-time dynamic exchange. In order to study this mechanism at the single-aggregate level, the super-resolution optical microscopy technique STORM is utilized.
FIGURE 1.19. Schematic model of BTA-fibers attached to the surface via linker molecules.
Modification of glass surfaces will be performed using silane chemistry. For anti-fouling behaviour, perfluorinated molecules will be used to create glass with a very high surface energy, and pegylated compounds will be used for their known anti-fouling characteristics in biomimetic anti-fouling coatings. After functionalization of the glass surfaces, the anti-fouling properties will be analysed using fluorescence microscopy and quantitative STORM measurements.
Subsequently, a commercially available silane linker molecule with an NHS-ester functional-ized end-group will be incorporated in the obtained anti-fouling surface for reactivation of this surface. Analysis using fluorescence microscopy will show if BTA-fibers indeed bind to these linker molecules, while maintaining their fiber-like shape and dynamic behaviour.
The final goal is to come up with a general method of producing glass coverslips that can be used for real-time imaging of the dynamics of supramolecular fibers in solution. If surface treatment is promising and successful, also other compounds and structures can be studied via selective binding to these glass coverslips.
C
HAPTER2
BTA-
SYNTHESIS AND DYE LABELINGL
ately, state-of-the-art experiments aim to investigate the dynamics of BTA-fibers using fluorescence microscopy, and for that it is necessary to acquire BTA-fibers with fluorescent properties. These are obtained by assembling fibers in which a small percentage of the monomers are labeled with a fluorescent dye molecule. This chapter describes how these BTA-molecules are obtained after reduction of BTA-triazide and subsequent coupling of BTA-triamine to a fluorescent cyanine dye (Cy3 and Cy5).2.1 Reduction of azide
A BTA-monomer has three arms with functional end-groups, schematically depicted in figure 2.1. The commercially available fluorescent dye molecule we use to couple with the BTA contains an NHS-ester reactive group. To react with our BTA, we need to obtain a BTA-monomer with primary amines at the ends of the three arms.
The triamine cannot be stored for too long, because it is unstable over time. The BTA-triazide does not degrade appreciably over time when stored at minus 20 degrees Celsius, and was therefore used as the starting material for the synthesis. Thus, the first reaction step is a reduction from precursor BTA-triazide to BTA-triamine. The reaction scheme of this reduction is shown in figure 2.2.
We used Staudinger conditions for the reduction that was performed. This reaction was named after Hermann Staudinger, who in 1919 reported that azides could react with triphenylphosphine to form phosphazide.[28] The reaction produces a high phosphazide yield and is nowadays commonly referred to as the Staudinger reduction. The exact reaction mechanism is shown in figure 2.3. It entails a nucleophilic attack of the phosphine at the terminal nitrogen atom of the azide, that eventually produces an iminophosphorane intermediate. Nitrogen gas is evolved
FIGURE2.1. Different end-groups of a BTA-monomer.
FIGURE2.2. Reaction scheme of reduction of BTA-triazide to BTA-triamine.
during this reaction, making it irreversible. In the second step, this intermediate is hydrolysed to form the amine and triphenylphosphine oxide. The reaction conditions are very mild and are here preferred over hydrogenation using palladium on carbon.
Exact experimental details can be found in the materials and methods section at the end of this chapter. Analysis using1H-NMR and IR-spectroscopy showed complete reduction of the starting material. The spectra can be found in appendix A. The desired product was obtained with a yield of 70 mg (55µmol), or 74%.
2.2. COUPLING WITH FLUORESCENT DYE
FIGURE2.3. Reaction mechanism of Staudinger reduction.
2.2 Coupling with fluorescent dye
Next step was coupling the obtained BTA-triamine with a fluorescent dye. Commercially available NHS-esters of the dye were available for this. In this case, two different dyes were used, namely the Cy3 and Cy5 cyanine dyes, because of their high extinction coefficient and their fluorophore emission maximum in the green and red region. These dyes have an activated ester that can react with primary amines.
The coupling of this dye with an amine is strongly pH-dependent reaction. At a low pH-value, the amino group is protonated, so that no reaction takes place. The optimal pH-value for reaction
The coupling of this dye with an amine is strongly pH-dependent reaction. At a low pH-value, the amino group is protonated, so that no reaction takes place. The optimal pH-value for reaction