I use a RIPA lysis buffer containing RNAseOUT 1:2000
I cross-link the cells with 1% formaldehyde to stabilize RNP complexes, but this has a disastrous effect on RNA quality (though on the odd time when the RNA quality is slightly better the IP is very specific). I believe the RNA quality is severely compromised during the cross-link reversal step: 45 min @ 70 degrees. The issue is that most of these experiments are not reproducible because the RNA quality varies so much which is reflected in the PCR. I would just like help finding a lysis buffer that promotes the stability of RNA while not being too denaturing such that the antibody is detroyed/ finding a way to stabilize the RNA during the cross-link reversal step.
Here's my experimental timeline
Day 0
1. Transfect cells
Day 1:
2. X-link cells (optional)
3. Collect cells
4. Add lysates to beads (keep 1% inputs)
5. Incubate 60 min @RT
6. Wash 5x 900 uL cold stringent buffer
7. Elute 1hr w/FLAG peptide
8. Reverse X-links 45min @ 70oC
9. Add Trizol LS (15 min)
10. Freeze @ -80oC
Day 3:
11. Extract RNA
12. Make cDNA
To me the problem is not the lysis buffer composition but the X-link step, formaldehyde has a disastrous effect on both DNA and RNA and requires to fine tune the X-link time to stabilize the RNA-Protein complex without damaging RNA too much, it can range from 2 minutes up to 30 minutes. Also the reversal time is quite important, you are using 45 minutes but I found protocols with up to 4 hours (see link).
http://munin.uit.no/bitstream/handle/10037/4847/thesis.pdf?sequence=2
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