I have designed and assembled a 10kb plasmid containing SbfI and NcoI restriction sites flanking a promoter region, which I am attempting to excise and replace with other promoters. When I do a double digest with both enzymes, I do not get my expected 750 bp band. I have already sequenced my plasmid and it definitely contains both of my restriction sites where I am expecting them.
I have digested my plasmid with SbfI and NcoI individually several times and it looks like each enzyme is working as I am seeing a linearised product on an agarose gel. Both enzymes are HF supplied by NEB and follow the same protocol with the same reaction conditions.
I've tried running the reactions for longer doing 30 min, 1 hr, 3 hr digests. I've used DNA from different (although Sbf is unstable after 1 hr) minipreps, different quantities of DNA (0.5 to 5 ug), different volumes of SbfI (0.5 to 3 ul), brand new restriction enzymes and rCutsmart buffers.
I've included a gel image below. Using a 1% agarose gel ran at 90V for 45 min and a Quick Load Plus DNA ladder.
I don't know what else to try or what else could possible be going wrong.
Any help would be greatly appreciated.
Dear Michael Cagney
Indeed, you tried all possible ways and designed excellent experimental setups. Yet, the following suggestions can be helpful:
1. Please check the enzyme units required for per ug DNA and add accordingly.
2. Since you are not getting 750bp band, may be double digestion is not working. You can still get a linear plasmid band in case of single digestion. To ensure, a reaction can be designed with 3 tubes- first one with SbfI only, second one with NcoI only and third one with both enzymes. Tube 1 and 2 can be of smaller reaction volume. If all three tubes show single band, then we can confirm, both enzymes are working.
3. It is recommended to aliquot buffer in small volumes to avoid repeated freezing and thawing.
4. Although it is HF enzyme, you can try a 100uL reaction volume with an overnight reaction time.
Hope it will work.
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