Hi everyone,

I work with the oospores (average diameter 27.7 µm) of a plant-pathogenic oomycete. In searching for antagonists of them, I incubate them with enzymes in solution or co-culture them with other microbes. I then assess viability changes using a combination of dyes - fluorescein diacetate and DITO-3 iodide.

My problem is that I always end up with a large amount of residual enzyme (and even buffer) or microbial residue in the tubes at the time of staining, which leads to many unexpected effects on the dyes (fluorescein diacetate in particular) and skewed results. I wash with sterile ultra-pure water and centrifugation twice before staining. I have tried adding more washes but it did not help and significantly increased the duration of the assay as well as increasing oospore loss (need to maintain oospore numbers for counting later on).

I have been thinking about filtration set-ups that might allow me to catch anything above, say, 15 µm and let through everything else. But I work with 24+ 1.5 ml tubes for each experiment, so the apparatus would need to work with this in mind and all the micro-centrifuge filtration options I can see are the typical 0.22/0.45 µm pore size (and would be a wasteful and expensive option anyway). Another limiting factor is the need to easily resuspend the oospores that I catch, in good numbers, to be stained. And this is a medium-throughput assay with small volumes, which also limits options.

I have also thought about flow cytometry/FACS, but the machines I have access to for these are on the other side of campus and would add an hr+ to an already long pipeline, where all I want is to reduce signal noise from residual enzymes. This option may be of more use down the line, when microbe-microbe interactions are a bigger focus.

Does anyone have any experience with this sort of very specific scenario? Or any ideas for how to more effectively wash my oospores clean before staining?