I am trying to detect DNA-protein covalent crosslinks which are induced by my drug treatment. I am following the RADAR assay protocol. Briefly, I am lysing cells using a buffer containing 6M GITC, EDTA, N-Lauryl Sarcosine and Triton X-100, followed by precipitation of the genome with 100% ethanol. The DNA is then washed twice with 70 % ethanol and then resuspended in 8mM NaOH. The DNA is then quantified by Nanodrop, and 500ng of each sample is loaded using a biorad slot blotting apparatus using vacuum. I am using a nitrocellulose membrane for blotting. Post blotting, the membrane is blocked with 2% milk for 1 hour, followed by probing. I am facing issue with reproducibility. I have tried shearing the DNA by sonication as well, without any improvement in results. There are gross sample-to-sample as well as experiment-to-experiment variations. Kindly suggest what can be done to improve the reliability. Do I need to bake the membrane or UV crosslink it?
I would try using less dna to ensure that all of the dna can bind and also bake it on to the membrane to minimise washing off. If you are resuspending in NaOH then positively charged nylon is probably a better immobilising membrane if it is available
Thank you. I did try with baking, and it significantly improved my results.
Unfortunately it's hard to achieve reproducibility with slot blots. I suggest you to load different DNA amounts and choose the 3 within a linearity curve of the signal. Then, always blot these 3 amounts per sample.
if possible, avoid ECL detection because it's not linear.
Alice Meroni Thank you for your response. I can try the idea of three amounts per sample. I have to detect the protein by Western Blotting after spotting the samples. I am currently detecting by ECL only. Is there any other method of detecting with similar sensitivity?
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