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no insert found - (Feb/21/2014 )

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I am trying to clone a 3.8Kb gene into 5.7Kb vector ( pFBDM )
Here is my protocol:

Vector preparation:
10ul of vector was diogested with two enzymes ( 1ul of XmaI and Kpn-I HF in CutSmart buffer , enzymes were from NEB as fresh stock ) at 37deg C for 3 hrs, then gel purify ( Qiagen gel elusion kit ).
Note: the DNA was not exposed to UV, I used another lane as standard and cut
at the corresponding site.

cDNA was obtained by reverse transcription ( NEB RNase - reverse transcriptasem, primed by oligo-dT )
4 extra bases are added for efficient digestion. According to NEB manual, it will be sifficient for 100% activity.
amplification: LongAmp Taq ( NEB ). I saw a sharp band on agarose gel.
The PCR product was purified prior to restriction enzymes digestion ( Qiagen kit )
Enzymes digestion: same as vector, I purify the DNA again after digestion with Qiagen kit.

50ng of digested vector was used in ligation
vector:insert ratio: 1:2
ligation 16 degC O/N ( NEB ligase )
negative control: vector + ligase only, no insert

negative control: 3 colonies
experimental plate: around 50 colonies

I picked 8 colonies and grew O/N at 37 deg C. Plasmid was purified using Qiagen kit. The plasmids preparations were digested with two enzymes, however, no insert found. If digested with single enzymes, the plasmid was of same size with original vector.

Thanks for your opinion.


When you cut with your two enzymes, how big was the piece you cut out (i.e. base pairs between the 2 restriction sites).  If it wasn't much (<100), chances are that when you were cutting your cut band out of your gel, you got a little uncut plasmid in the mix and that's what you're seeing on the experimental plate.  


I included a negative control ( cut vector only , no insert ), only 3 colonies yielded. The negative control essentially excluded the possibility.


I'd like to know more about yoru reverse transcriptase reaction. You said you used an oligo dT primer -- did it have the XmaI or KpnI cut site? Or is there another primer used for the gene you are interested in. It sounds as if you have done many things correctly, but I'm uncertain about your final vector and insert cooncentration. I'd recommend next determining if your insert cut ends ligate efficiently. Do this by taking your PCR product, cutting with only one of the two enzymes, and ligating. You should get a double length fragment. Do this for both enzymes. You could also get rid of background by making your vector with PCR, which then allows you to digest the template with DpnI. I'm a little concerned about your transformation efficiency, because I would actually assume your background should be much higher after vector digestion and gel purification.


I am using the oligo-dT primer not gene specific primer. I did not include RE in my RT primer.


I used 50ng of cut vector for ligation, insert/vector ratio was 2:1. The insert is 3.8Kb long; vector is 5.7Kb long.


The transformation efficiency of my competent cells is 10^8 CFU/ug plasmid. I tested it in positive control and worked fine.

Negative control yielded few colonies suggesting that self ligation is minimal; I am curious about why experimental plate contained so many background colonies?

Is it possible that bacteria delete the insert?


Thanks again.


So, are you amplifying the insert with a different primer after the RT reaction? Where does the restriction site for your insert come from?


You quote a transformation efficiency number, but is that hoped-for, or measured? The ability to transform with prepared plasmid tells you essentially nothing about the required efficiency, unless it is done at the 10 pg of plasmid level.


For future reference, it is a bad idea to post to two fora. The conversation happening in the other forum is relevant and should be in one place. I disagree with pernesseblue about ligation concentration. Too high a vector and insert concentration leads to concatamers, which fail to transform. 20-50 ng of vector in a ligation is the sweet spot, along with equimolar insert.

Are you using quick ligation mix, or the normal ligation buffer?


Oligo dT primer was used for reverse transcription. I did not use gene specific primer in RT.

I tested the transformation efficiency with 10pg of pUC19.

I used the regular T4 ligase and set up 16 deg C O/N ligation.




So, I don't understand how you expect to clone a fragment which does not have a restriction site at one end into a KpnI/XmaI digested vector. One end of  your insert wil have a poly T region, which won't clone.


The RE sites are included in my PCR primer. So my PCR product contain the RE sites. The PCR product and vector were both digested with Kpn1 and Xma1.


So, after RT reaction with an oligo dT primer, you do a PCR on the cDNA with a pair of primers, both of which have a restriction site. And this is what you see on a gel. Is that right?

If so, I repeat my suggestion about testing the ligation efficiency of single-enzyme cut versions of this PCR product.

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