I’m having trouble with my Western blots and wanted to see if anyone has experienced something similar.
I’m running 4–12% or 12–20% Bis-Tris gels (NuPAGE / Bolt), loading ~30–50 µg of protein from cultured cells. For transfer, I’m using the Invitrogen iBlot 3 Dry Blotting System, typically running the default 7–8-minute “Mixed MW Proteins” program with PVDF membranes. Blocking, antibody incubations, and washes all follow standard protocols (5% milk or BSA in TBST, ~1 hour block, overnight primary at 4°C, etc.).
The problem: when I image the blot—even the molecular weight ladder shows absolutely no signal. I’m using a modern dual-color prestained ladder (e.g., SeeBlue Plus2) to visually monitor transfer. The ladder transfers cleanly to the membrane, but when I develop using ECL reagents on a digital imager, there are no chemiluminescent bands at all—no ladder, no endogenous controls, nothing.
Could proteins be transferring through the membrane during the iBlot run, leaving me with an empty membrane?