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Problem with cDNA synthesis


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#1 shwetaT

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Posted 31 July 2018 - 07:55 AM

Hello,

 

I am new to doing RT-PCR. I have done  a bit of standardization. But stuck with one problem which I dont understand where could the problem possibly be....

 

I use Trizol (TaKaRa) for RNA isolation followed by cDNA synthesis (verso kit, Thermo).

 

one problem I am facing is DNA contamination for which I have reduced the original culture for RNA isolation so as to get as little DNA as possible. Then I treat the sample with DNAse (37 degree C for 15 mins) followed by LiCl precipitation (7M) and three volumes of Ethanol for 3 hrs at -20 degree C.

 

For cDNA synthesis, I have tried different temperatures : 42 degree C for 1 hour and 50 degree C and 55 degree C for 30 minutes

 

The whole cycle for cDNA synthesis:

RNA (200-400ng) random hexamers and water at 65 degree C for 5 mins, add other reagents and incubation at 42/ 50/ 55 degree C for ihr/ 30 minutes, followed by 94 degree C for 2 mins.

 

when I do RT-PCR using cDNA, there is no difference in Ct values between NRTC control and experimental set when house-keeping genes gyrB and 16S are used. The melt curves seem to be okay

 

I have done verification of these primers with PCR and I get single sharp band of expected size.

 

Can anyone help please?? Is there a problem with cDNA synthesis....what could be the possible reason for no difference in the Ct values??

 

Thanks

 

 

 



#2 OldCloner

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Posted 01 August 2018 - 10:32 AM

First, can we assume your No Template (NTC) control was negative, meaning you don’t have a general lab/reagent contamination problem?

 

Next, I assume the CT values that do not differ are low Cts, meaning you are getting a lot of amplification for both the cDNA and RNA (no RT control).  And this is for two commonly expressed bacterial housekeeping genes, so you are extracting RNA from bacteria?  The RNA you extract will contain a load of ribosomal RNA including 16S (it is not translated into protein), right? Whereas the gyrB is a message that would be translated into protein. Hmm.

 

The problem with qPCR of bacterial samples is that you can’t design primers to cross exon-exon splice boundaries- at least not for most bacteria. So gDNA contamination could be a big problem. So, how would one test for the presence of bacterial gDNA specifically? Maybe you could try designing primers to amplify a part of the genome that is not expressed in one transcript. Put one primer in one operon and the other in a nearby but different operon (This assumes you are using a well-characterized bacterium where this sort of information is available in the literature). If you can amplify this gDNA fragment from your RNA (before reverse transcription) then that would indicate that yes, you do have a problem getting rid of the gDNA, and you need to work on that.

You can also try another kit for isolating the RNA in the first place. Dunno if that would help or not.



#3 merlav

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Posted 01 August 2018 - 11:54 AM

Run RNA samples in a denaturant or native gel or a automated electrophoresis before doing the RT, if DNA is present do another digestion with DNase and run it in another gel.  Once your RNA is clean from DNA you perform the RT making sure that you prepare an RT-(no enzyme) for each sample, then perform an end point PCR with a HKG.  You should get one strong band in the RT+ and in the RT- no bands (sometimes you may see primer dimers).  Then you perform the QPCR.  Did you perform an optimization for the primer concentration and sample quantity for the QPCR??? If not, that could be one of the problems of the Ct values.  


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