University of Groningen
Novel fluorescent probes and analysis methods for single-molecule and single-cell
microscopy
Smit, Jochem
DOI:10.33612/diss.102269758
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Publication date: 2019
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Smit, J. (2019). Novel fluorescent probes and analysis methods for single-molecule and single-cell microscopy. Rijksuniversiteit Groningen. https://doi.org/10.33612/diss.102269758
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“It is very easy to answer many [..] fundamental biological questions; you just look at the thing!”
Richard Feynmann, 1959
In his famous talk ‘There’s plenty of room at the bottom’, Richard Feynman indi-cated to his audience the emergence of a new field of physics: a field we would now refer to as ‘nanotechnology’. During the lecture, he analysed the problems and possibilities of manipulating and controlling things on a small scale. By showing that, when shrinking letters down to the atomic level, it is possible to write the entire Encyclopaedia
Britan-nica on a tiny speck of dust, he demonstrated that there is indeed plenty of room at
the bottom.
Feynman’s goal in exploring the atomic scale was not to produce pocket-sized books with infinite information; but instead in the possibilities of transforming the contem-porary room-sized computers into miniaturized versions with orders of magnitude more computational power while reducing power consumption. He suggested that, through the development of ‘tiny machines’, we might be able to manipulate atoms and perform chemical synthesis via mechanical manipulation. As of today, Feynman’s vision has largely become reality. Computers scaled down to nanometre dimensions scale can be found in every person’s pocket. Artificial ‘molecular machines’, although realized by chemists rather than physicists, represent a rapidly emerging development1–3.
However, molecular machines have been around for as long as biological life itself. In fact motor proteins located in every living cell are the first ‘tiny machines’. They are responsible for every process occurring in cells of all life domains; generate energy, create new proteins, recycle old proteins, copy genetic information, regulate intracellular concentrations of metabolites and ions, and act as proofreading devices or as clocks. The tiny machines able to manipulate atoms catalysing chemical synthesis are abundant and central in nature; they are commonly called ‘enzymes’.
Over the course of the last few decades, the view of the chemical biology of the cell shifted from simple chemical kinetics where compounds A and B come together in a stochastic and diffusion-controlled encounter leading to the formation of a product4,5,
to a view of tightly controlled and orchestrated machineries consisting of many subunits
and components, working together in a highly coordinated manner leading to directional and polarized motions and conformational changes. Every life process is based on such a fundamental thermodynamic principle: molecular motors dissipate free energy which is transformed into mechanical work, thereby allowing biological life to exist out-of-equilibrium.
But how can we elucidate the fundamental working mechanisms of molecular motors at the cornerstone of molecular biology? Certainly, Richard Feynman was right about one thing: you just look at the thing! However, when it comes to looking at things on the molecular scale, is it not very easy. Proteins and their complexes are ranging from sizes of a few nanometre to several tens of nanometres thus to be able to ‘look at things’ on this scale, powerful microscopes are needed.
Back in 1959, answering these questions was virtually impossible. The most pow-erful electron microscopes were unable to resolve biological structures due to technical limitations. Therefore, Feynman was calling for a 100-fold improvement of the electron microscope’s resolution. Although 100-fold resolution increase has not been reached, resolving protein structures at atomic resolution has been achieved, in large part though the development of cryogenic electron microscopy6–8. This technique, specifically
single-particle cryo-EM, has given rise to a revolution in structural biology, leading to the structural determination of many protein machineries (Figure 1.1).
However, biological processes do not take place in a vacuum or at cryogenic tempera-tures. In contrast; such processes are warm, fuzzy, heterogeneous and, most importantly, dynamic. If we want to figure out the molecular details of these machineries, we need to study them as close as possible to the physiological conditions. Therefore, the tool of choice needs to be able to resolve dynamics at the molecular level, be sensitive to static and dynamic heterogeneity and be able to do so in a minimally invasive manner. Optical fluorescence microscopy is a tool that fulfills the above-mentioned requirements.
Optical Microscopy
One of the main challenges of optical microscopy is achieving the required optical contrast needed for visualization of biological structures. Biological samples are typically transparent with a refractive index close to that of water, therefore the absorption and scattering processes which generate contrast are minimal. The red onion, a unique biological sample able to provide a high contrast image without any sample preparation
a b c d
Figure 1.1 | Examples of protein-based molecular motors. a) Adenosine triphosphate (ATP) synthase9. This motor consists of two rotating parts, generating ATP that “fuels” many other biological processes. b) RNA polymerase10. Composed of a dozen different proteins, RNA polymerase is responsible for reading DNA and copying the genetic code into RNA. c) Ribosome subunits11. The ribosome is one of the largest biological machines composed of proteins as well as RNA. Both subunits dictate proteins synthesis based on the instructions by reading processively messenger RNA. d) Pilus machine12. Another example of a rotary motor, the pilus is thought to extend and shorten via rotation of the motor. Protein structures are elucidated by Cryo-EM or X-ray crystallography and were obtained from the protein data bank13and illustrated by David Goodsell.
is the reason for being adopted by teachers all over the world gratefully in order to introduce students to the wondrous world of optical microscopy (Figure 1.2).
Figure 1.2 | Red onion cells under an optical microscope. Image: r/oddlysatisfying u/Jamesthebutler18
Several different variations on the normal transmitted light microscopy setup have been developed over the years to enhance contrast. In phase contrast microscopy14,15,
the phase of the light waves is used to enhance the contrast. Light scattered by the
biological sample is phase-shifted by 90 degrees and the illumination light is passed through an annulus and subsequently, after the objective, a phase shift ring. This leads to constructive interference between the sample-light and the background light, increas-ing contrast. The phase contrast microscope allowed scientist to study cell division and the cellular structure of living cells for the first time.
Fluorescence Microscopy
Instead of using scattering or absorption for contrast, the fluorescence microscope uses the physical phenomenon of fluorescence to generate contrast. The principle relies on the labelling of the sample with fluorescent probes. These probes can absorb photons of a given wavelength and then emit a photon with a longer wavelength. This shift in wavelength or photon energy is called the Stokes shift16, and based on this principle
the fluorescence microscope can generate the ultimate contrast.
By using mirrors and filters which transmit or reflect only light of a specific wave-length window, the high intensity light used to excite fluorophores can be effectively separated from the fluorescent emission of the fluorescent probes used (Figure 1.3). These dichroic filters or interference filters can attenuate the excitation light with a factor in excess of 106 while transmitting most of the fluorescent light. This allows the observation of fluorescent labels on a dark background.
Single molecules
The combination of this high contrast technique together with improvement in detector sensitivity and background-reducing illumination techniques, have enabled fluorescence microscopes to detect single fluorescent molecules. First demonstrated in 197617, this
breakthrough has pushed fluorescence microscopy to the absolute limit of sensitivity and has opened a whole new field of biophysical research. In biological systems, pro-cesses take place far from equilibrium, often with many distinct interconverting states or conformational dynamics. This temporal heterogeneity is often combined with spatial heterogeneity – cell-to-cell variations, differences in local chemical environment or the presence of chemically distinct subpopulations.
These kind of heterogeneities in biological samples cannot be unravelled by standard ensemble techniques. A typical ensemble experiment probes a mixture of 1016molecules
Figure 1.3 | A schematic representation of a fluorescence microscopy setup. Excitation light is coupled into the objective via a dichroic beam-splitter to illuminate the sample. The emitted fluorescent light is red-shifted with respect to the excitation light and is collected onto a charge coupled device (CCD) camera. Image: Principles of Fluorescence Spectroscopy, Lakowicz
simultaneously. Without synchronization, which is not feasible in in vivo experiments, any heterogeneity in the sample will be averaged out over the population.
Diffraction limit
Although through fluorescence the contrast can be pushed to the limit of a single emitter, the use of optical waves for detection sets a fundamental limit on the maximally achievable spatial resolution. This limit is a consequence of the wave nature of light: diffraction occurs when it encounters obstacles or lenses. Due to this ‘bending of the waves’, no features smaller than roughly twice the wavelength of the light used can be resolved. This fundamental principle is known as Abbe’s diffraction limit18. For optical microscopes, this places the limit around 200 nm (Figure 1.4), which prevents scientists from exploring the ‘nanoscale’ of biology by optical microscopy.
Optical Super-Resolution
Although Abbe’s diffraction limit is literally written in stone, in 2014 the Nobel prize in chemistry was awarded to Willian E. Moerner, Eric Betzig and Stefan Hell for breaking
Figure 1.4 | The different length scales found in biology. Abbe’s diffraction limit prevents optical microscopes to resolve biological structures below 200 nm. Image: Johan Jarnestad (The Royal Swedish Academy of Sciences)
exactly this limit. They developed a new set of techniques now referred to as ‘super-resolution microscopy’ or ‘nanoscopy’ as they go beyond classical ‘super-resolution limits and take microscopy into the nano-domain.
The super-resolution principles can be divided into two categories: targeted readout and stochastic readout19–23. In both, the resolution limit is bypassed by extending the imaging into the temporal dimension. Under normal fluorescence microscopy condi-tions, fluorophores in the same diffraction-limited region are excited together and emit together, making it impossible to discern them individually. In super-resolution mi-croscopy, fluorophores are switched between ‘on’ and ‘off’ states, allowing individual emitters to be separated in time. In the targeted readout technique Stimulated Emis-sion Depletion (STED)24,25, alongside the excitation light, a second doughnut-shaped beam of light is applied. This second beam induces off-switching in the fluorophores at defined coordinates, whereas only the fluorophores in the middle intensity minimum of the doughnut is allowed to fluoresce.
The methods Photo-Activated Localization Microscopy (PALM)26,27and Stochastic
Optical Reconstruction Microscopy (STORM)28 (collectively single-molecule
localiza-tion microscopy (SMLM)) stochastically switch all fluorophores in the field-of-view using imaging buffers with chemical additives or photochemical on-switching, in order to sep-arate the emission temporally. The location of the individual emitters can then be pinpointed and its exact location can be determined with high precision.
Typical resolutions obtained are in the 20-50 nm range, and theoretically the resolu-tion is only limited by the number of photons that can be collected from a single fluo-rophore. Using cryogenic temperatures, where photochemical processes are slowed down
and more photons can be collected per fluorophore, resolutions down to 2 Angstrom have been reported29.
Fluorescent labels
Not only the maximum attainable spatial resolution depends critically on the fluorescent label used. The temporal resolution depends on the fluorescent label as well – no dynamics faster than the photon emission rate can be observed. Moreover, the chemical conjugation and labelling capabilities of the label determine which cellular structures or biological systems can be investigated. The chemical and photophysical properties of the fluorescent label are therefore one of the most important parameters in a fluorescent microscopy experiment.
The two most commonly used classes of fluorescent labels are organic fluorophores and fluorescent proteins. Both have their unique advantages and disadvantages, mak-ing each complementary, suitable for specific applications. Organic fluorophores are designed and synthesized by organic chemists. Thus, all tools of the chemistry toolbox are available to tailor a fluorophore to specific demands. Therefore, these probes fre-quently exhibit desirable photophysical properties, such as high extinction coefficients, high quantum yields and stable fluorescence emission. Their absorption and fluores-cence maxima can be tuned across the visible spectrum and a plethora of chemical conjugation options are available. These can be biorthogonal chemical conjugations, such as copper-free click chemistry30,31, or bioinspired conjugation via SNAP32,33- or
Halo34-tags.
Fluorescent proteins, on the other hand, have the major advantage of being geneti-cally encodable. Through standard molecular biology procedures, the genetic sequence of a fluorescent protein can be attached to a gene encoding for a protein of interest. Then, when the cell produces the chimeric fusion, the fluorescent protein “labels” the target protein. This is a significant advantage over synthetic organic fluorophores which need to be inserted into the cell externally and conjugated to the biological target through carefully designed conjugation strategies. The chromophore in fluorescent pro-teins is formed via a cyclization reaction between several amino acids in the core of a the beta barrel of the fluorescent protein. This limits the structural freedom of fluorescent protein chromophore compared to synthetic organic fluorophores, and as a result the photophysical performance of fluorescent proteins often suffers from low brightness and
fluctuations in fluorescence emission.
Despite the fact that fluorescent protein design is limited by linking the 21 different naturally occurring amino acids in sequence, many different fluorescent proteins have been developed and discovered with a wide range of varying properties. The first flu-orescent protein to be isolated was the green fluflu-orescent protein from A. Victoria35,36, which has its main peak of absorption in the ultraviolet37. Today, the available
fluores-cent proteins span the whole visible spectrum, and variants include photoactivatable or photochromic proteins38–40.
The basics of fluorescence
The electronic structure of a fluorophore molecule dictates the basic principles of fluores-cence. The energy levels of fluorophores are distinct electronic states which correspond to molecular orbitals of the molecule. These molecular orbitals are a result of a com-bination of the atomic orbitals within the molecule and the energy spacing between the highest occupied molecular orbital (HOMO) and the lowest unoccupied molecular orbital (LUMO) is what determines the colour and thus the absorption spectrum of the fluorophore.
In fluorescent organic molecules, the HOMO usually corresponds to the S0 energy
level (Figure 1.5). When the electrons occupy this state, the molecules are said to be in the ground state. From this state, a molecule can absorb a photon with an energy matching the transition gap with a probability roughly proportional to the absorption cross section of the molecule. Usually, the molecule is excited to a higher vibrational energy level of the excited singlet state. In fluorescence microscopy the molecules are excited to the energy level S1. Within a timescale of picoseconds, the molecules relax
to the lowest vibrational level of the S1 state. From this state the molecule can emit
a photon as fluorescent light (within several nanoseconds) and the molecule returns to S0, again in a higher vibrational state which rapidly decays as heat via a process called
internal conversion.
The loss of energy via vibrational relaxation has as a consequence that the fluorescent photon emitted has a lower energy and therefore a longer wavelength than the photon absorbed by the molecule. This shift in energy is called the Stokes shift.
The cycling betweenS0andS1electronic states is an idealization of the true
Figure 1.5 | Jablonski diagram depicting the electronic states of fluorescent molecules and their interconversion processes. Image: Principles of Fluorescence Spectroscopy, Lakowicz.
emission, including internal conversion (vibrational coupling), collisional quenching, en-ergy transfer, photoinduced chemical reactions and intersystem crossing.
Intersystem crossing is the transition of one electronic state to another with different spin multiplicity. This type of transition is formally forbidden in non-relativistic quantum theory, but through spin-orbit coupling all organic fluorophores have a small probability of transitioning from the first excited singlet state to the first excited triplet state.
The Triplet state and consequences for photobleaching
The same quantum mechanical rules dictate that also the transition back from T1 to
S0 is quantum mechanically forbidden. Therefore triplet state lifetimes are long-lived,
up to a hundred microseconds41 or milliseconds42. This fact, together with the high
reactivity of the unpaired electrons of the triplet state, makes the triplet-state pathway the main contributor of photochemical destruction of the fluorophore. This process is called photobleaching and leads to immediate loss of fluorescent signal. It is exactly this photochemical process that is currently the limiting factor for fluorescence microscopy studies.
The role of oxygen
Molecular oxygen has a very peculiar electronic structure: its ground state is twofold degenerate, and according to Hund’s rule of maximizing spin it follows that the electronic
ground state is a triplet state. In a typical imaging buffer oxygen is present in the sub-millimolar range and it has a high propensity to react with radical ion species formed after triplet state formation (Type I photosensitization) or directly with the fluorophore triplet state43 (Type II photosensitization)44. The result of this process is a variety of
reactive oxygen species (ROS), including superoxide anions (O-2), singlet oxygen (1O2)
and hydroxyl radicals (•OH). The reaction of these species with the fluorophores can lead to irreversible photobleaching45,46.
This thesis aims to resolve some of the technical hurdles faced in fluorescence mi-croscopy. In the first part, we expand on the previously established concept of self-healing fluorophores to enhance their photostability. The second part focusses on the imaging of bacterial cells, which includes software for data analysis of fluorescence mi-croscopy datasets followed by the development of a fluorescent probe and a clinical application of fluorescence imaging.
A brief summary of the chapters is presented below.
Chapter 2
Organic fluorophores are augmented with self-healing properties by conjugating them to triplet-state quenchers. Specifically, their performances in different types of optical super-resolution microscopy is investigated. We show that the conjugates give superior performance under STED conditions and high excitation energies. For STORM, we find that the photo-switching kinetics are strongly influenced by conjugated to a triplet-state quencher.
Chapter 3
The mechanisms of intramolecular triplet-state quenching is investigated in more detail. In competition experiments between intramolecular and intermolecular experiments we investigate the relative rates of triplet state quenching. It is reported that although intramolecular triplet-state quenching is capable of quenching the triplet state rapidly, the presence of alternative bleaching pathways limits the obtained photo stability.
Chapter 4
The software package ‘ColiCoords’, developed for the analysis of fluorescence microscopy data collected for rod-shaped bacterial cells is presented. The software is released as an open-source analysis platform written in the programming language Python. The main
feature is to transform Cartesian coordinates, either pixel coordinates or single molecule localization microscopy (SMLM) data, to cellular coordinates. This transformation allows for data collected from many different cells to be aligned and combined, as well as the projection of fluorescent signal along any axis, be it radial, longitudinal, or angular.
Chapter 5
Organic fluorophores conjugated to aminoglycoside antibiotics are applied as a probe to selectively image bacteria. We find that the probes selectively stain the gram-negative bacteria E. coli, P. aeruginosa and K. oxytoca whereas the gram-positive bacteria S.
aureus, S. epidermidis and L. lactis are not stained by the probe. We show the probe
NeomycinB-RhodamineB can be used in antimicrobial photodynamic therapy. Further-more, when administered intravenously in mouse models, the probes enable the detection and selective imaging of gram-negative infections.
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