I use commercially available Qdot secondary antibodies that work in general. However, compared with e.g. Alexa488 labeling in the same sample I do not get the same specific staining, i.e. actin stress fibers are barely visible. The staining looks a bit spotty and there is not a very high degree of colocalization between Qdot/Alexa.
I tested a range of fixation/permeabilization methods (Acetone, methanol, formaldehyde, TritonX100 hi/lo), because I assume it is an accessibility problem. This is what the LifeTech Support suggested and what I somehow saw in a more or less good microtubule staining - there, at the thickest part of the cell, the microtubule Qdot staining was weaker compared to Alexa. I also tested embedding media (Cytoseal 60 vs. Mowiol) and dehydration before embedding. Everything without success so far...
Any other ideas or working protocols?
Thanks in advance!
Hi Bernd, this difference could be due to the actin antibody specificities.
If both the Alexa and QD antibodies are binding to a multiple sites in actin, then you will get a stronger signal with the Alexa labeled antibody.
Because of the QD size, there will be more steric hindrance of QD antibody binding to actin compared to Alexa labeled antibodies.
Probably some useful points will be in attached article.
Article Comparative study of Surface Plasmon Resonance, electrochemi...
Hi, I am not an expert in QD staining method. But, in my experience I found fixation in paraformaldehyde is crucial for preserving the ultrastructure of actin cytoskeleton. Fluorescent dye-conjugated phalloidin is very efficient in staining actin cytoskeleton. However, if you have to use QD antibodies, you may want check the filter sets you are using. I have read that use of the conventional filter sets used for imaging dye-stained samples are less efficient for QD stained samples.
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