.. My Rat spinal cord samples are preserved in sucrose 30% .. I cut the sections using cryostat at 40 micron, All my buffer solutions for 1ry & 2ry Antibodies, blocking solution, washes are composed of TBS 10 x 10 % , BSA 0.25 % , 0.3 % Triton X - 100 that I use all through my protocol.
N.B, Blocking Solution (which is always Horse Serum) not matching that of 2ry antibodies' Hosts.
Problem: I have my staining all through my samples not confined to certain cells as supposed or layers of spinal cord. I have used different antibodies using this protocol and the results are the same, unspecific stains everywhere .. Can it be a result of using Triton 0.3 % all through the whole steps? Thanks a lot.
Rinse in distilled water, shorten the time and reduce the concentration.
I have many questions for you to think about:
How is the tissue fixed before cryopreservation? How do you freeze the tissue? How do you mount the sildes? How does a secondary-only-control look like? What is the pH of your buffers? 40µm is quite thick, do you really need it that thick?
Triton X-100 is quite strong, try lower concentrations ot milder detergents. Tween20 is a typical one.
I also agree with the others, that a higher concentration of BSA (2-5%) might help. Is there a reason why you use TBS and not PBS buffer? If you want to look for phosphates be aware that BSA is not the best choice.
The suggestions presented here are correct. If you cut 10 -15 micron frozen sections simply reducing the thickness will reduce background. Also , since you are using sectioned frozen material, do you really need to extract it? Usually whole cells need to be extracted to allow for AB penetration. If you do need to extract then do it once, probably for only 2 min at RT, after mounting the sections prior to AB exposure. You can also try changing the slide coating, if you are using albumin fixative. Good luck.
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