Mounting

One thing that people tend to forget is that the most important (and hardest) part of doing microscopy is mounting the specimen. You can have the best stain ever because you spent weeks optimizing some protocol and it's all useless if you screw up during mounting. Two things are important:

1) You want to preserve the three-dimensional structure of the tissue as well as possible. This is especially true if you want to do morphometrics from 3D reconstructions, but also for simple maximum projections you want to have the tissue to look as close as possible to what it's like in situ. To achieve that, you want to make sure that you don't do any damage mechanically (see below), and you want to choose the right mounting medium. It has been described that potentially any histochemistry can screw up the structure of the specimen. At least in the case of arthropod wholemount preparations, the only real problem seems to be the mounting medium. Mounting in glycerol is a bad idea. It creates torsions and all kinds of anisometric shrinkage arifacts. The agent of choice in my opinion is methyl salicylate (wintergreen oil). If you dehydrate carefully (30,50,70,90,100% EtOH) and then let the tissue sit in a 50:50 EtOH:methyl salicylate for an hour or 2 before mounting in pure methyl salicylate, all you get is a little isometric shrinking (see Bucher et al (2000). Correction methods for three-dimensional reconstructions from confocal images: I. tissue shrinking and axial scaling. J Neurosci Methods 100:135-143).

2) You want to make sure that you can use the best possible objective. Higher resolution objectives have fairly short working distances and you don't want to end up not being able to use one of those just because your cover slip is a mile away from the tissue. In theory, most objectives are optimized for spherical corrections for a 170micron cover slip and if you deviate from that you will lose signal. But that is only important if you have a really, really weak signal. Usually you want a thinner cover slip to win some working distance. Most companies have cover slips in different thickness ranges. I recommend ESCO No. 0, which reliably seems to be 100 microns thick. It's nice to have a range of different ones around, because you can also use them to keep the cover slip from touching the specimen (see figure).

Mounting the STG:

Here's what I do. I leave the nerves (stn, dvn, avn, agn) fairly long and pin the ganglion into a thinly coated sylgard dish. After fixing, but before dehydrating (in buffer), I cut the sylgard around the ganglion with a scalpel but leave everything in place. After dehydration, I take the cut out piece of sylgard out of the 100% EtOH and transfer it to a small glas vial filled with the methyl salicylate/EtOH mixture. I leave that in a vaseline-sealed dry glass container (with silica gel on the bottom), to keep the EtOH from drawing water. After 1-2h, I transfer the sylgard piece to a sylgard coated glass dish and pin it down. I put a drop of pure methyl salicylate on top. Now I cut the nerves proximal to the pins that hold them. Then I carefully place a shard of coverslip underneath the ganglion and use that as a "gurney" to transfer the ganglion to the slide without having it fall dry and without coiling any nerves. I usually glue 2 coverslips on a microscope slide with nailpolish from the top. The STG goes into the gap between them. I carefully position everything the way I want it and then fill up with mounting medium, remove the cover slip shard, put a cover slip on top, and seal everything with nail polish. Make sure it's clear nail polish, otherwise the color will dissolve into the medium. Using 100micron thick coverslip let's me even use the 100x oil objective we have.