I've got 4 highly similar nucleotide fragment and i wish to design specific primers for each of them. Their protein coding are about ~80% similar. I tried to use blastn (NCBI): align 2 sequences (megablast, discontinuous megablast, balstn) and pasted in 2 of my sequences. And the result was: no similarity for both megablast, and ~70% similarity for blastn from 112 to 253. The thing is, by looking directly at 2 sequences, the similarity started from 1st base, not base 112! Did i somehow altered the setting or what happened? How should i align more than 2 sequences together so that i can have a clear look on them?
I use the Turbo DNA free kit von ambion. Works well and is easy. Basically you add DNAse I and Buffer, incubate for 30 min at 37°C then you add their bead based inactivation reagent, incubate, spin and take the supernatant. For me it work 80 - 90% of the time.
I have used Turbo DNA free kit (Ambion) as well. In our samples Turbo DNA free kit degraded about half of the amount of the DNA contamination in the samples. There was still some DNA left in all my RNA samples after DNA degradation
I've had the same issue. Used DNaseI from Sigma originally, then tried Turbo from Ambion. Turbo worked much much better than natural DNase I, but still left detectable levels of DNA. Then I read a paper somewhere that the use of the RNeasy on-column digestion coupled with Turbo got nearly all bacterial DNA out, but that a new product from Epicentre would do the same, but without having to do the on-column. It's called Baseline Zero. We've been using that since, but of course it still leaves some DNA contamination.
I've come to the conclusion purely through assumption that there is a certain concentration of DNA at which the DNase is no longer able to digest. There's still a little DNA, but too little for digestion... at least at a reasonable rate.
So I ended up digesting a high concentration of RNA (and therefore fairly high DNA concentration) with Baseline Zero, then diluted the DNase-treated RNA down. I figured the digestion would drop the DNA level down as far as it would go, then the dilution would drop it even further.
I think I would digest 2 ug of total RNA in a 20 ul reaction (so 100 ng/ul concentration of RNA). Then I would dilute the RNA down to 10 ng/ul for use in first strand synthesis. For the first strand synthesis reaction (20 ul) I would use 1 ul of the 10 ng/ul RNA. So I was diluting the contaminant DNA (after DNase treatment) 10x after DNase treatment, then 20x going into the RT reaction for a total of 200x dilution of the "lowest" DNA level possible by DNase treatment.
At each step, I ran a PCR to determine if detectable DNA was present. Without DNase treatment, there was a very strong product. After DNase treatment, there was a weaker, but still strong product. After the 10x dilution, there was a very faint product, and in an RT-negative first strand synthesis reaction, there was NO product.
Also, I'd like to add, this is all done on bacterial RNA, so I don't have the luxury of spanning an intron, so DNA elimination is very important. Everything I've read regarding DNA in RNA samples is that there is virtually NO WAY to get rid of it, you just need to get it down to a manageable level, i.e., a concentration at which it is not detectable.