Hi, everyone,
I am optimizing my protocol of influenza PR8 plaque assay. I am following the classic protocol. But I ran into problems when I am trying to visualize plaques at the high virus dilution. My protocol is as below.
1. seed 5x10^5/well MDCK in 12-well plate and culture overnight (MEM medium with 5% FBS, 1% P/S, 1% Amp)
2. wash with MEM twice. infect MDCK with PR8 in infection medium (MEM medium with 0.1% BSA) and incubate for 1 hr at 37 degree. Rock the plate every 10 min.
3. wash with MEM once. lay 1% agar gel about 42-45 degree. (2xMEM with 7.5% BSA, P/S, Amp, NaHCO3, TPCK typsin mixed with 2% agar gel). Wait 10-15 min till the gel solidate.
4. Put the plate back to the incubator and incubate for 48hr.
5. Fix for 1 hr at RT with 4% formaldehyde. Remove gel and stain with 0.5% crystal blue for 5 min while shaking.
The result is shown in the attached picture. I could only see plaques when the virus concentration is high. I also tried 1:5 serial dilution. It still has the same problem. I tried to extend the culture from 48 hr to 72 hr. It did not help since the cell layer was fragile and washed off before crystal violet staining. Instead of visible plaques, I always saw the black dots in the middle virus dilution. The black dot is a cluster of dying cells under the microscope. So I am wondering how I could visualize the plaques at this dilution. Should I try low concentrated formaldehyde? shorter fixation (10-20 min)? put the plate in 4 degree fridge overnight instead of fixation? use lower concentrated crystal violet (0.1-0.2%)? seed fewer MDCK to make a monolayer instead of a thicker cell layer?
I know PR8 is not a good plaque former. Could anyone let me know if you ran into the similar issue and how to solve the problem?
Many thanks!