Which approach to staining do you take? A bath, sitting drop? How is your sample attached to the slide? Do you have some adhesive or is it one of the electrostatic slides? If you're experiencing liquid bead formation that would suggest local hydrophobicities, we need to figure out where it came from to work around it. 

Samples are attached electrostatically. DAPI is mixed with the mounting medium and then coverslipped as a final step after rehydration. I don't think it is some immiscibility with the PBS, like residual clearing agent or anything like that. All sections are fully de-waxed and rehydrated, and fluorescence in all other channels is strong and well resolved under the scope without artifact. My impression is that water is being caught by the texture of the section, in between these roughly 200µm ridges that constitute the biofilm surface morphology. I spoke with a colleague from Barnard and she said that she has noticed this little local dilution of DAPI as well, but just images around it - choosing sections that don;t carry the artifact. I can also do that. It isn't so pervasive that it is in all of my sections, but I'd ideally like it if it were in none. 

OK, I wonder if the way you apply dapi might be the problem. I was always taught that dapi must be rinsed off after staining and that, contrary to what the books say, dapi does fluoresce not intercalated into DNA. I know that some people use dapi in mounting media, but I've also seen a comparison of HeLas washed and unwashed from excess dapi and they really are not the same. I can imagine a situation where this would cause problems especially in the case of a biofilm containing some eDNA.

Is there a chance you could do a 5 min staining in PBS and them wash it of with some more PBS before you mount the slide? I know it's a pain, been there, did that, but it might be worth the effort if it gives you the image you want!

You have to consider that the salts in the PBS might cause some problems with the dapi, we use dapi (1:1000) by leaving the cover slip directly on it for 15 mins then wash it 5 times (10 mins) with PBS. But we always dip it in ddw prior to mounting so the salts from the PBS will be remove. We also gently dry the some of the water by touching with the tip of the cover slip with a piece of paper. If you do consider a wash in a well, try gently fill it with water through the edges of the well and not directly on the sample, it helps (I also have a problem with the staining and the wash like you, but this works for me most of the time without washing away the samples)

The salts are a good point. I have noted the exothermic precipitate that forms between the buffer and the ethanol. I've always wondered if the osmolarity of my final rehydration wash was an issue post fixation, and I've considered rehydrating to water instead of PBS. Of course pH needs to be considered. Thank you for your help gentlemen. I have some things to try now. I'll start by optimizing the wash step, like Adam suggests, and I'll also try dipping in di-H2O prior to mounting in the anti-fade. I'll let you know how it goes. 

No problem, Good Luck!