Oh, my protocol is pretty standard as well. Using 40um frozen sections, I wash 3 x 20 min. with PBS, then permeablize for 1 hour with 0.3% Tx-100 in PBS (This step might not actually be necessary since the blocking and Ab buffers have 0.3% Tx-100 as well, but I guess I'm a little superstitious about deviating from a working protocol). I block using 5-10% normal animal serum of the species in which my secondary Ab was produced, diluted in PBS-Tx (block for 2-4 hours depending on what else is going on that day). I have had background issues as well when using BSA, although I know some people use it routinely with good results. For antibody buffers, I use 3% animal serum in PBS-Tx- there is usually some leftover blocking buffer that I can just dilute down. I do incubation overnight at 4C then wash 3x the next day, and then my secondary for 1-1.5hr at room temp. Wash again 3x then I mount with a paintbrush. Of course incubation times and conditions may vary with Ab. But this general protocol seems to work for most things.
For section handling, I use the netwells (12 well format) and just accept that I'm going to lose some sections to damage. I don't have too many problems with tissue damage in P21 or older mouse spinal cord sections, but I tried some P7 tissue recently and they had much more damage from the netwells. So if your tissue is fragile then maybe use the hook or paintbrush method to transfer. With a paintbrush you can probably pick up a bunch at once so it saves some time. Alternatively, Corning seems to have a wide variety of different well inserts made of different materials and meshes so maybe it's just a matter of finding the right product for fragile tissue. This is going to sound weird, but I know our lab uses pantyhose (like pieces of the material, not the whole garment) to hold and support live spinal cord slices while they're incubating prior to ephys recording, and I've often wondered if I could build some small holders our of this material for IHC use.