I am, strangely, having trouble detecting actin in muscle lysates. Tissues are lysed in RIPA buffer and run under reduced conditions.
I typically load approximately 40ug in my Western and then perform a semidry transfer. This is confirmed by coomassie and ponceau staining. I block the membrane with 5% NGS in PBS-T(0.05%) for 1 hour RT. Primary antibodies (mouse monoclonal anti-beta actin) are 1:10,000 in 2% NGS PBS-T(0.05%) and incubate for 1 hour RT. 3 x 15 min washes in PBS-T(0.05%). Secondaries (IRDye 800CW Goat anti-Mouse IgG H+L) are made up 1:10,000 in 2% NGS PBS-T(0.05%) and are incubated in the dark for 1 hour RT. 2 x 15 min washes in PBS-T(0.05%) and 1 x 15 min wash in PBS are performed.
The beta actin band should be extremely strong in muscle. When I did this using CHO cell lysates I got an OK actin band. But in muscle lysates (both in mine and other students') I get a Western as attached. Any help would be appreciated.
You may incubate over night at 4 degree with primary antibody. I think your primary antibody concentration is too diluted. You may use TBST and skim dry milk powder for blocking instead of PBST for washing. Best of luck
Do you work on mouse tissue? If that is, try to find another abs not being a mouse origin.
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