I have been attempting to isolate lymphocytes from mouse lymph nodes (inguinal and axillary) with varying and limited success. I have tried to different methods.

1. Place LNs into ice cold PBS. In a 100 mm tissue culture dish containing approximately 10 ml PBS, disrupt LNs by rubbing between frosted surface of two autoclaved slides. Wash slide surfaces off and strain through a 70 micron cell strainer into a 50 ml conical. Mash any large pieces with the back end of a 25 ml syringe plunger. Rinse strainer with an additional 10 ml PBS. Centrifuge for 7.5 minutes @ 400xg. Resuspend in 1 ml RPMI and count with trypan blue.

2. Place LNs into ice cold RPMI. Place cell strainer in a 60 mm tissue culture dish containing 2 ml RPMI such that the strainer portion is submerged. Place the LNs into the strainer and gently mash with the back of a 2 ml syringe plunger. Count cells with trypan blue.

With the first method I typically obtain anywhere from 1-4 x 10^ cells total with 50-70% viability.

With the second method its and order of magnitude less with 1% viability, but in this second method there are approximately 10^7 cells that are 99% dead.

I know that my yield should be in the 10^7 range and I am not sure if PBS v RPMI is the critical factor or the aggressiveness with which I am disrupting the LNs.

Any help would be greatly appreciated!