I generally use nonimmune Rabbit Serum or the Rabbit IgG from Vector 
Labs diluted to the same protein concentration as my primary antibody 
as my Negative controls.  The researcher I work for wants to use 
adsorption controls for the negative control now.  We have been able 
to obtain the immunogen from the company who produced the polyclonal 
antibody.  I have tried several times and keep getting a lot of 
backgroung staining - seems like everything is sticking everywhere.  
I titrated the primary antibody out - to get best signal to noise 
ratio.  Using that concentration (dilution) then added the immunogen 
at a 5 times more concentration to the primary antibody and allowed 
to incubate for 2 hour @ RT, then diluted to working concentration.  
I used that (primary + immunogen) inplace of the primary antibody.  I 
keep getting all sorts of staining with this control.  My "negative", 
with the Rabbit IgG is clean, the "positive" with the primary 
antibody is staining where it should.  The researcher INSISTS this 
should work; however, I just do not know where to go from here.  I 
understand the theory behind everything.  I know the adsorption 
controls work well in western blots; however, has any one tried them 
as negative controls for tissue sections?

Primary antibody:  Rabbit anti-insulin 
Blocking peptide:  Peptide used as immunogen
Detection Kit:  Vector Rabbit IgG ABC Elite Kit
Chromogen:  DAB
Tissue:  Mice pancreas

Beforehand, Thanks for any comments or suggestions.

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