I cloned the PCR product (gel checked) into the p-GEM T Easy vector system and extracted plasmid DNA. Now I am doing restriction enzyme digestion to check if the target got successfully inserted. I used EcoRI in the Buffer H system with 200 ng DNA as recommended, incubated at 37 C for 1 h, loaded 8 uL of samples with 2 uL of 5x loading dye, and ran the 1.2% gel (target ~200 bp) at 65-70 V for ~1.2 h. However, the result showed that the target bands were there, but with very low intensity compared to the backbone. And what also confused me was that the control without enzyme added, which I expected to see a single plasmid band close to or beyond my sample backbone bands, showed up at a lower position than the backbones with some other unknown bands (as shown in the third lane from right in the picture). So my questions are:
1) Why my target bands had such low intensity? Does it mean an incomplete digestion (I used both 30 min and 1 h incubation but observed no significant change in intensity)? If so, should I increase the incubation time or the enzymatic activity my deteriorated under -20 storage over time?
2) Why the no-enzyme control showed multiple bands and the plasmid band was shown at a lower position than the backbone bands in comparison with the samples with enzymes?
I greatly appreciate your kind help and suggestions!
Hi Yang Xiao, I presumed your desired band was located at a lower position than your desired position, which in this case, it meant your targeted band was smaller in size compared to the desired band. There could be many reasons, for example you ran the gel for too long and led to the further down migration of the band, or could be due to further digestion of the DNA which led to smaller band, wrong band, contamination, non-specific target etc... Many reasons for this, which you might need to troubleshoot..
However, when we mentioned about the band intensity, it usually referred to the "brightness" of a band, which might indicate DNA abundancy and low intensity might be related to DNA degradation etc..
All the best!
Hi Chong, thank you very much for your answer. You are exactly right that my target band should be at a lower position than the band shown in the picture. If running gel for too long is the problem and leading to the further migration of the band, then why the band showing up was at a higher position, shouldn't it be at a lower position then? Also if the DNA was over digested, shouldn't the band also be at a lower position? If my DNA was contaminated or degraded by nuclease, then why the plasmid DNA was not affected?
I greatly appreciate your kind reply and looking forward to your further suggestions. Thanks!
Hi there,
The detection of DNA by intercalant is quantitative if the vector is 10 times bigger than the insert then the signal detected for the vector will be 10 times more intense than for the insert. What is the size ratio between vector and insert? if the ratio is actually 10 then digesting totally 200ng of construct will result in 182ng of vector and 18ng of insert. The ratio of band intensity is relevant of the size ratio between vector and insert.
Hi Dominique, thank you very much for your kind response! My insert is about 200 bp while the backbone is about 3000 bp, so that is more than 10 times. Does it mean that seeing such target bands with low intensity is a normal result according to my insert : backbone ratio? And to get a brighter target band, I should use more plasmid DNA? But I still do not understand why my target bands were a bit higher than the expected position and the control without enzyme was located even lower than those being digested. Do you have any suggestions of improvements that I could make? Or do you think according to the ratio and the result, my plasmids are safe for sequencing?
Thank you so much for your generous help!
Hi again,
Non digested plasmid is actually a mix of different forms (there are 3 main forms : supercoiled, linear and relaxed(nicked)). The one running faster than linear is the supercoiled form so it's normal to see a band more mobile than the linear plasmid in the undigested sample. About the mobility of the insert it might be an effect due to the content of the digest mix reaction. To make an accurate determination of the band size you should actually prepare the ladder sample exactly the same as the plasmid digest sample (minus the restriction enzyme).
For more info about plasmid pattern on agarose gel:
https://biology.stackexchange.com/questions/19114/why-isolated-recombinant-plasmid-dna-without-any-restriction-is-often-detected
... about sequencing: just go for it : it's OK using miniprep samples.
Your gel picture looks perfect. The undigested vector is supercoiled so it will run faster than the linearized vector with only a small fragment (200 bp) released. Further, your target fragment (200 bp) will not be very bright if you used just 200 ng Plasmid DNA for digestion. I would suggest to use upto 2 micro gram plasmid DNA for digestion if you need more amount of the target fragment. If you are just using for ligation confirmation then 200 ng is fine, you can see the band released. Further, sequencing will confirm the identity of the cloned fragment. Also when small fragments (200 bp) are run for longer time on gel, the intensity of the bands get reduced so a bit shorter run time will improve the intensity of the released fragment.
I agree with Dominique and Kafeel. Your gel image is correct.
1) When you digest your DNA, smaller fragments show up less dense.
2) An undigested plasmid as a circular DNA runs always lower compared to the linear form of the plasmid.
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