5.1 Conventional Synapses
In this work I demonstrated that STED microscopy has the power of dissecting single synaptic vesicles in vivo using video-rate imaging. The resolution of the vesicles was limited to approximately 60 nm by the STED laser power, increasing the resolution by a factor of around 18 compared to confocal imaging. To further increase the resolution one could use higher STED laser power, but this would in turn substantially enhance photobleaching, which was already the case with the used settings. In addition, higher STED laser power would cause problems in collecting enough emitted photons from the reduced effective focal spot, and consequently, in separating the moving vesicles from background noise. Related to this, the fluorescent dyes and laser technologies need to be improved in parallel to reliably enhance resolution. Moreover, STED lasers with continuous wave beams would excite the fluorophores continuously, and thus allow for capturing more photons. Consequently, imaging of weaker signals, as well as imaging/ scanning with higher speeds over the sample would be possible. During this work it has been shown that STED with continuous wave beams can be applied to video-rate imaging on conventional synapses (Lauterbach et al., 2010).
However, as the resolution of the microscopes will presumably at some point achieve the size of proteins and antibodies, labeling methods need to be revisited. A gigantic primary and fluorescently-tagged secondary antibody complex used in immunofluorescence imaging would hinder the precise colocalization of two in principle exactly colocalizing proteins as their fluorophores could be separated some tens of nanometers.
In addition, protein colocalization studies would also need two-color STED imaging, which was already demonstrated for fixed cells, but awaits its application on living samples (Donnert et al., 2007). The application of a multi-color STED microscope with the ability of video-rate imaging would allow the investigation of the two vesicle pools simultaneously and would by far expand the here presented work. Questions one could address with a video-rate
1. What can we learn on the synaptic vesicle mobility when the two different pools (such as the recycling and resting vesicles) are investigated in parallel?
2. How do the vesicles move in relation to cellular elements like the AZ, is their movement more directed towards the AZ upon (strong) stimulation?
Video-rate STED imaging used in this work was only performed on cultured hippocampal neurons. As already mentioned STED microscopy is difficult to use in tissue, as it has a low penetration depth. However, it would be of great interest to have nanoscale resolution available for deep tissue imaging to allow vesicle mobility studies in native brain slices. One recent study used STED two-photon laser scanning microscopy to investigate spine morphology deep inside acute brain slices (Ding et al., 2009). By applying a pulsed two-photon excitation laser and a one-two-photon STED continuous wave laser they achieved at an imaging depth of around 100 µm a 3-fold increase in resolution compared to confocal imaging. Nevertheless, this resolution enhancement would not be sufficient for vesicle mobility studies as presented here.
A great advantage would be nanoscale resolution in all three dimensions in vivo. Three-dimensional STED imaging was already performed on fixed cultured cells by the implementation of a second STED laser, the z-doughnut. The microscope allowed nanoscale imaging of immunostained neurofilaments with a resolution of 45 x 45 x 108 nm (Wildanger et al., 2009). However, capturing a single image frame (10 m2, pixel size 10 nm) takes 4 minutes – which is unfeasible for fast in vivo investigations as most biological processes are very short lived.
Moreover, three-dimensional optical sectioning was applied to image the endoplasmatic reticulum inside a living cell, but not at high frame-rates (Hein et al., 2008).
Another form of three-dimensional nanoscale imaging can be achieved with an isoSTED microscope. This technique relies on the 4Pi microscope system (Hell and Stelzer, 1992). A 4Pi microscope is equipped with two opposing objective lenses that focus into the same focal area. The molecules in the focal spot are illuminated by both lenses, which also collect the emitted fluorescence. Consequently, the interference of the two opposing light beams results in the resolution increase along the optical z-axis of about 5-7 fold (Bewersdorf et al., 2006).
Since this method only increases the z-axis resolution one is still left with a poor lateral resolution. Thus, Schmidt and colleagues combined STED microscopy with 4Pi microscopy
(Schmidt et al., 2008). The focal spot in the resulting isoSTED microscope thus has x,y,z-resolution of around 40-50 nm. Three-dimensional nanoscopy was only once applied to single synaptic vesicle investigations of fixed hippocampal cultured neurons (Wilhelm et al., 2010), and is awaiting its application on living specimens.
5.2 Sensory Inner Hair Cells
The here presented results of membrane recycling in IHCs were achieved with high-resolution electron microscopy of FM dye labeled and photo-oxidized preparations. The investigations were based on morphological observations of the recycling organelles to uncover the vesicle retrieval pathway on the single organelle level. Since vesicles in hair cells cannot be live-labeled without difficulties with FM dyes or antibodies as used for conventional synapses, vesicle recycling studies on IHCs in vivo are challenging (FM dyes (Gaffield and Betz, 2006), antibodies (Kraszewski et al., 1995)). However, electron microscopy allowed me to perform the first morphological identification of the endocytic pathway in these cells.
Nevertheless, the data presented here needs to be strengthened by defining the properties of the large cisternae and their vesicle budding mechanism. The next steps are therefore aimed at investigating the protein complement of the recently endocytosed organelles (cisternae, organelles at the apex and bottom). Related to this, correlative microscopy could be used, in which the IHCs are stained by a fluorescent fixable marker (e.g. dextran, see Results) and immunolabeled against proteins of interest. The preparations will then be embedded in a plastic matrix, which can be processed in 50-80 nm thin-sections before imaging in high-resolution STED mode and subsequently with electron microscopy (see for example Grabenbauer et al., 2005; Watanabe et al., 2010). This would allow to characterize simultaneously both the morphology and the protein identity of the recycling organelles.
Moreover, this technique would also allow to perform three-dimensional nanoscopy on fixed IHCs with a resolution of less than 80 nm in all directions (Punge et al., 2008).
In general the questions one should address are:
1. Which organelles contain synaptic vesicle proteins (and are thus directly involved in
2. Are proteins of the clathrin machinery (clathrin, AP2, amphiphysin) involved in vesicle recycling (at cisternae as well as at the other recycling organelles)?
3. Is dynamin used for the fission step of the recycling organelles?
4. Are Golgi complex or endoplasmic reticulum markers present on the recycling organelles?
5. Do recycling organelles contain endosome markers?
Although high-resolution fluorescence microscopy techniques (such as STED) are not applicable to study deep inside tissues STED imaging was performed on fixed IHCs with less than 100 nm resolution at a tissue depth between 15 to 25 µm (Frank et al., 2010). A great advantage also for IHC studies would be three-dimensional optical sectioning at nanoscale resolution, as the dimensions of an IHC are much larger compared to that of a small conventional synapse. Finally, after finding the proper vesicle marker the application to in vivo studies would be optimal to precisely localize the vesicle and traffic them inside the IHC.
Recently, a pH-sensitive green fluorescent protein (GFP)-based sensors, termed pHluorin, was introduced which allows the optical monitoring of vesicle cycling (exo-/ endocytosis) during synaptic activity (Miesenböck et al., 1998). When the pH-sensitive GFP is tagged to a protein domain in the synaptic vesicle lumen its GFP-fluorescence is quenched by the acidic environment inside the vesicle (pH 5.6). In contrast to the vesicle lumen the extracellular space has a neutral pH at around 7.4. Vesicles that fuse with the plasma membrane during stimulation expose their lumen to the extracellular space (to a neutral pH) and consequently the pHluorin fluorescence has a higher quantum yield compared to the quenched situation.
Upon endocytosis the protein gets incorporated into the lumen of the vesicle, where the environment becomes acidic from the activity of a vesicular proton pump (ATPase) and suppresses its fluorescence, hence the vesicles become dark again. While the so far generated constructs of the general synaptic vesicle proteins synaptophysin (pHluorin construct: sypHy (Granseth et al., 2006)), synaptobrevin-2 (synapto-pHluorin (Miesenböck et al., 1998)), the vesicular glutamate transporter-1 (vGlut-pHluorin (Voglmaier et al., 2006)) and synaptotagmin 1 (pHluoro-tagmin (Fernández-Alfonso et al., 2006)) are all absent from IHCs, new pHluorin variants need to be specifically generated for sensory inner hair cells.
The expression of a pHluorin-tagged protein in IHCs would allow in vivo-studies on vesicle recycling to analyze exo-/ endocytosis kinetics in IHCs, as well as the mobility of fused vesicles and the site of their retrieval.