Although our study (Kamin et al., 2014) represents a step forward towards the understanding of IHC physiology, further investigations would require the association of recycling organelles with synaptic vesicle markers in a high-resolution microscopy technique. That is the main aim of the project described in this thesis. Transmission electron microscopy (TEM) is the technique that has reached the highest resolution so far (<2 nm in biological samples) (Faas et al., 2012). However, immunolabeling techniques suitable for EM microscopy require laborious procedures that at the end offer poor epitope recognition and low labeling density. Therefore, a fluorescence microscopy technique would be more convenient for the easy sample preparation and high imaging throughput.
A major drawback of light-based microscopy techniques is the diffraction of light when passing through the lenses of a microscope. The German physicist Ernst Abbe postulated (1873) that a beam of light with a wavelength λ, converging to a lens of refractive index n and aperture angle θ, will produce a focal spot with a full width at half maximum (FWHM) given by the formula ∆r = λ/2(nSinθ), where nSinθ is equivalent to what is nowadays known as the objective numerical aperture (NA) (Hell, 2007). This means that two point sources of light imaged with a conventional fluorescence microscope cannot be told apart if they are closer than approximately 200 to 300 nm, for emission wavelengths in the range of the visual spectrum. In the case of synaptic vesicles and trafficking organelles, typically in the size range of 30 nm to a few hundreds of nm, and densely packed in the cytoplasmic volume,
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this is an important difficulty.
In 1994 Stefan Hell and Jan Wichman postulated the principles of what would become the first far-field microscopy technique overcoming Abbe’s diffraction barrier: stimulated emission depletion (STED) microscopy (Figure 1.6) (Hell and Wichmann, 1994). Six years later, the first practical demonstration was published, imaging the plasma membrane of Saccharomyces cerevisiae and Escherichia coli cells (Klar et al., 2000). STED microscopy is based on sample laser scanning, as done in confocal microscopy. However, in this technique an excitation laser is spatially overlapped with a second depletion laser. The front wave of the latter is physically modified with a vortex plate so at to generate a toroid or doughnut shape with zero intensity in the center and maximum intensity at the borders. The wavelength of the depletion laser is chosen to match the red-side tail of the fluorophore’s emission spectrum. In the doughnut center, where only the excitation laser is present, fluorophores are allowed to emit photons spontaneously. In contrast, in the outer area were both lasers are present, fluorophores will reach the excited state but the depletion beam will induce them to emit photons at its same red-shifted wavelength. These photons (stimulated emission) can be filtered out to only detect those produced at the doughnut center, resulting in a narrower diffraction-unlimited point spread function (PSF) (Klar et al., 2000; Hell et al., 2004). Abbe’s equation is therefore redefined as∆r≈
λ
/(2NA 1+I/IS ), where I is the maximum intensity of the depletion (STED) beam, and Is corresponds to the saturation intensity required to reduce fluorescence probability by half. Thereby, higher depletion laser intensities will render narrower imaged areas, increasing the resolution of the microscope (Nägerl et al., 2008; Moneron et al., 2010).In the past, STED microscopy has fostered important findings related to constitutive trafficking pathways and synaptic function: it was useful to establish the molecular players and steps required for cargo sorting and budding in early endosomes (Barysch, 2009). It validated the role of endosomal sorting in the segregation of plasma membrane proteins from recycled synaptic vesicles (Hoopmann et al., 2010). It revealed that only 40-50% of the SNARE proteins Syntaxin 1 and SNAP-25 are located at putative vesicle release sites, while the rest dispersed on the plasma membrane (Punge et al., 2008). It also helped to establish that synaptotagmin 1 remains clustered on the plasma membrane upon synaptic vesicle exocytosis (Willig et al., 2006). Among other studies, it also helped to put forward a novel hypothesis: the majority of synaptic vesicles found in the synaptic terminal (>80%) do not participate in neurotransmitter release, but rather act as a buffer for proteins involved in
29 vesicle recycling, keeping them concentrated for their eventual use (Denker et al., 2011b).
Using a commercial STED microscopy setup, like the one I used in this study, resolution can go down to 30-40 nm in biological preparations. The relatively short image acquisition time and the instant delivery of diffraction-unlimited images, without the need for further signal computation, are the main advantages of STED microscopy over other high-resolution microscopy techniques.
Figure 1.6 Working principle of high-resolution STED microscopy.
A. Schematic representation of the basic elements in a STED microscope. An excitation beam (green) is spatially overlapped with a depletion beam (red) and scanned over the sample as in conventional confocal microscopy. The depletion beam is modified by a phase modulator to create a zero intensity region in its center, resulting in a toroid or doughnut-shaped front wave.
Furthermore, the wavelength of the depletion beam is selected to fall in the red side tail of the fluorophore’s emission spectrum. In the sample, the fluorophores located at the center of the depletion doughnut are excited and allowed to emit photons spontaneously (yellow), which are collected by the detection device. In contrast, the excited fluorophores at the borders of the depletion doughnut are stimulated to emit photons in the red-shifted wavelength of the depletion beam, which is filtered out of the detection range. Hence, photons are only collected from a smaller subdiffraction-sized area. B. Example images of a neuronal soma and process imaged in confocal (left) and STED microscopy (right). Membrane labeling was performed with the novel endocytosis marker developed in this study (see section 3.2.3). Note that intracellular organelles are only distinguishable in the improved STED image. Scale bar, 2 μm.
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