This seems like perhaps the wrong place to ask this, but maybe asking on a site directed towards school children would give me a more clear, basic answer. This is a laboratory technique question.

I am a molecular biology tech, as of 2 weeks ago. I have spent less than ten days doing these techniques, and I am concerned about one thing in particular that I'd like to "nip in the bud" as soon as I can. After doing a few minipreps and subsequent restriction digests, my gels have shown what looks like gradually more and more supercoiled and nicked plasmid, gradually less linearized plasmid, and none of the desired insert.

I was wondering if high amounts of supercoiled and nicked plasmid is a sign of bad technique during the miniprep or is it a problem from something previous/upstream like the ligation or the transformation?

Thank you,

Frustrated Tech

PS. I have attached pictures of some of my gels, in chronological order. The dates are in the file name. As you can see, they get uglier and uglier. Note: the negative control is the first sample on the bottom row next to the ladder/size marker.

Post's pictures

digestion8-11-11.jpg, 13.49 kb, 406 x 506

restdigFIV-8-25-11_2.jpg, 21.75 kb, 293 x 381

restrictiondig8-12-11-2.jpg, 13.35 kb, 329 x 431

I wouldn't worry about the ligation/transformation as being the source of your problems. If you were able to get colonies on a selective plate, your plasmid is intact. Are the sample shown freshly generated plasmids, or are they the same sample? It looks like you might have some nucleases contaminating your minipreps, which could explain the multiple bands in the negative of your latest gel. If your restriction digests are also not working properly, that strongly suggests a DNA purity problem.

Without details of your methods, I can't really give you any specifics on what might be going wrong, but check your miniprep protocol and ask some more experienced lab-mates for some help.

I can suggest a few general things;

Check that the strain of cell you are using is suitable for DNA preparation. For example, using endA+ ecoli (like JM101, TG1, BL21) requires an additional wash step when using a miniprep kit. If you are using a kit, the most likely source of contamination is inefficient washing of the column.

If you are doing phenol/chloroform extractions, make sure that the phase interface is completely clear before moving to the next step. The presence of non-polar solvents will disrupt the activity of the restriction enzymes, so make sure your DNA pellet is completely dry before re-dissolving it.

If you are adding an RNAse (one that was not provided in a kit), heat it at 96 degrees  C for a few minutes to kill off any residual DNAses.

Good luck!

I wouldn't be disheartened, most of us have had digests like yours before! To add to John's general list, try and be as gentle as possible in the initial lysis step - this can help reduce nicking. If you are doing a precipitation with something like isopropanol or even EtOH make sure you give the DNA miniprep pellet a good wash (even a few minutes) in 70% EtOH before resuspending your sample. Pay particular attention to John's point about removing excess non-polar solvents.  If you are like me you may be prone to adding too much restriction enzyme (with the accompanying buffer such as glycerol which may inhibit cutting) on occasions, so you may be able to use much less enzyme, especially for 'easy' cutters like BamHI, EcoRI or HindIII.

The problem seems to be that the restriction digestions are not going to completion.  For reference, 1 U restriction enzyme can digest 1 ug of DNA in a 50 ul reaction in 1 hour.  My guess is that these reactions have too much DNA and/or too little enzyme and/or they've not been incubated for enough time.  This is suggested by the presence of the undigested negative control band(s) in the lanes where digestion clearly occurred.  

I would recommend routinely overdigesting samples; to do this, calculate how many units of restriction enzyme are needed to completely digest the DNA in 1h, then use a 5-10x excess -- this will help compensate for differences in DNA concentration and purity.  The total enzyme in the reaction however must never exceed 10% v/v to avoid inhibition by glycerol (this is the maximum upper limit, but aim for 5% v/v).  The exception to this is when the restriction enzyme has reported star activity, in which case overdigestion should be avoided (though such enzymes are relatively uncommon).

This should work as far as cleaning up the restriction digests goes; if not then DNA cleanup should help.  You can also consider colony PCR for screening of putative clones (the prefered method in our lab for initial screening due to improved speed and cost).

As John says, it's a bit difficult to know exactly what is going on here without some more details. Rough amounts of DNA cut (e.g., what % of DNA from what volume and type of bacterial culture), enzyme(s) used and how many units over what time, etc, etc? Most of the common enzymes cut very quickly (and often to completion) and an hour digestion is usually more than enough. There is (or appears to be) some 'partialing' in the top gel (bottom half). I have had results similar to the middle gel (which looks the 'worst') and changing DNA concentration, enzyme units/time/etc has made no difference. Assuming that the a recommended standard amount of DNA marker has been loaded, and looking at the 'shape' of the DNA bands, it appears that much less than 1ug of DNA was in each digest. I agree with John that on the info provided a DNA purity problem (and consistency between preps) is the most likely problem.

Last edited by Steve Lolait (28th Aug 2011 09:56:04)

Yes Steve I agree with you and John that impurities in the DNA may explain the results.  NEB recommends overdigesting with 5-10x RE to compensate for this and other variables -- if it still doesn't work then cleaning up the DNA should work.

OP - as mentioned above, some extra info. on the bacteria, the miniprep protocol, and the restriction digest protocol would help us to narrow down the possibilities.