A sample is embedded in agar, glued to a slide, mounted in a water bath, and placed under the microscope. A vibratome associated with the microscope exposes the top of the sample block. The microscope performs a tile scan over the exposed sample surface then slices a thin section off the block. The microscope alternates between tile scanning and imaging until the whole sample has been acquired. Physical sections are taken approximately every 50 microns and optical sectioning is possible to allow for finer resolution in z. The slices sink down to the bottom of the water bath and are usually discarded once acquisition is complete.
Any sample that can be embedded in agar, cut on a vibratome, and contains a fluorescent signal can be imaged. Fixed tissue exhibits rich autofluorescence so label-free structural imaging is possible. Imaging of a wide range of fluorophores is, of course, also possible. The sample should ideally be brightly labeled to allow for short integration times and therefore faster imaging. Typical use cases that work well include counting labeled somata, mapping dye-labeled electrode tracks in brains, tracing bulk fibre projections in brains. Higher resolution scenarios, such as brain-wide tracing of individual axons is possible but very challenging.
Datasets created by automated microscopy techniques can potentially be very large and so will usually require automated analysis techniques in order to extract meaning. It becomes particularly important to select imaging resolutions that are appropriate for the question at hand otherwise dataset size can quickly enter the TB range and acquisitions stretch into multiple days. Parameters such as voxel size and integration time should be carefully chosen to be adequate for the downstream analysis pipeline and not over-sampled. Often a quality that is perfectly adequate seems noisy or low resolution to those more familiar with confocal microscopy or even automated slide scanners.