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Need good fixation protocol for visualizing microtubules in cultured cells by el

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#1 Lyanesse



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Posted 08 February 2010 - 01:59 PM

Hello all,

I am trying to find a good fixation protocol for cultured cells (neuronal progenitor cells in this case) that preserves cytoskeletal/microtubule structure for electron microscopy. I have previously prepared cells for EM with PFA and got mediocre results, which I now understand is because PFA is not optimal for preserving microtubules.

When fixing cells for immunostaining, I get beautiful preservation of microtubules following this protocol for glutaraldehyde fixation of the tubulin cytoskeleton: http://mitchison.med...ocols/gen1.html . Basically, this consists of a 30sec extraction (0.5% TritonX) step in microtubule stabilizing buffer, followed by fixation for 10 mins in 0.5% glutaraldehyde at 37C. Then sodium borohydride wash to quench the free glutaraldehyde.

Would this protocol work for EM if I just followed it with postfixation in OsO4, or would the detergent extraction cause artifacts in the EM?

Also, if I can use this protocol for EM, is the sodium borohydride step necessary? I'm not planning to do immuno-EM.

Or are there other protocols that anyone has experience with for this application? I also tried preparing cells according to Deitch and Banker (J Neurosci 1993), where you add 3.5% gluta directly to the culture medium and incubate for 15 mins at 37C, followed by postfixation in 1%OsO4. Even after just the glutaraldehyde fixation step the cells looked all bunched up and somewhat fragmented, so it was not too surprising that the EM did not look very good (fragmented cells, nuclei bunched up all together).

Any suggestions would be greatly appreciated! Thank you!


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