I’ve cultured mouse ESCs and differentiated into neurons. I proceed with PFA 1% fixation, cell lysis using lysis buffer containing 50mM Tris-HCl pH=7, 5mM EDTA, 1% SDS and inhibition cocktail (PMSF, proteinase inhibitors and RNASin), sonication (3 cycles, 6cycles and 12 cycles) and then RNA extraction. When I quantify I get good amount of RNA (in average 700ng/ul) but when I run a gel I can’t see bands. I used a control of total RNA and that one is ok and I see the two RNA bands and in the 3 samples (3 cycles, 6cycles and 12 cycles sonication) I see absolutely nothing!
I’ve denaturated the rna before running, I’ve already tried using TAE instead of TBE because TAE I used with lower pH 7,3.
please help me!
RNases are omnipresent enzymes. I am speculating that your target RNAs might have been degraded by RNAses. It is always good to perform RNA work as much as possible under RNases-free conditions.
Fixation with PFA will cross-link anything within 2angstroms so your RNA could be crosslinked to protein and not separating well in the trizol extraction - it may be staying in the organic phase. I would ask why you are fixing the cells and if it is necessary. PFA fixation can be reversed and likely needs to be to extract the RNA efficiently.
Please clarify the gel electrophoresis and detection methods you are using (e.g. Agarose or Polyacrylamide, what buffer, what voltage, what time, SYBR Gold, etc) that are not producing a band, and let us know what you might have that could be used as a positive control (e.g. an RNA prep that has previously produced a visible band). You may inadvertently be using a poor choice for these factors.
If you want a non-electrophoresis method, you could run RNA integrity analysis using an Aglient 2100. I believe your institution has access to one, but not be fluent in Italian I cannot confirm this suspicion (link below).
http://www.telegrid.enea.it/NuoviProgetti/Prisam_Grid_1621/Far/Prog%20Esecutivo%20Parallelomics.pdf
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