Hello,
	I'm relatively new to immunohistochemistry, and am trying to optimize 
a protocol for staining whole ant brains.  The brains are relatively 
small pieces of tissue (about 300 microns x 300 microns) that should be 
(if I understand correctly) well within the size range of tissue pieces 
that can be successfully processed whole.  I have been using the 
standard 4% paraformaldehyde in PBS at 4 C fix, and have been 
incubating in primary antibody 1:50 for 3 days at RT, and secondary 
1:200 for 2 days at RT.  Despite these long incubations I'm having 
trouble with antibody penetration, and am starting to look into 
permeabliziation methods.  I see several techniques in the literature: 
enzyme digestion, adding Triton-X 100 and/or DMSO to the washing and 
incubation buffers, and incubations in acetone or methanol after 
fixation, but there's no explanation (that I can find) about how these 
techniques work.  Does anyone have experience with these issues?  What 
does DMSO or Triton-X or acetone "do" to the tissues?  What kinds of 
concentrations have you found useful or not useful?  There is some 
indication of a protective, very very thin "sheath" of tissue around 
the actual neurons of the brain.  If this is the case, is enzymatic 
digestion of the tissue the only option?  Pros or cons of the different 
treatments?

I know this is a long and detailed question.  Thanks so much for any 
help you might provide.

Mario Muscedere       
Biology Department
Boston University
5 Cummington Street
Boston, MA, 02215 
Office: 409 BRB
Office #: 617-353-6977
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