My insert (3.7kb) concentration is 448ng/ul. In the double digestion reaction, I am using final concentration of insert of 2ug in a 50ul reaction setup. After incubating with restriction enzymes (EcoR1-HF and Sal1-HF) and running it on 1% agarose gel an extremely faint band is observed (the size was correct) - which cannot be used for ligation purposes. What optimization do I have to do to get a fair-intense band so that cloning purpose can be served?
Details:
Insert - 4.4ul
10XNEBuffer - 5ul
EcoR1 - 1ul
Sal1 - 1ul
38.6ul nuclease free water is added to make the volume upto 50ul.
Incubate at 370C for 3hrs.
Is it a PCR amplicon that you're using for digestion? or you're trying to cut out the insert from a construct?
Either way faint band doesn't correspond to the amount of DNA you're using i.e 2ug. Either you're using less amount of EtBr in the agarose gel or using very less exposure during imaging.
If you're digesting PCR amplicon you can directly do PCR cleanup after digestion. No need to run on the gel again. As this will reduce your recovery.
If your plasmid is large and if the total OD260 gives 448ng/ul then the actual insert concentration is much less than expected by the ratio of the insert size to the plasmid plus insert size. Also run an uncut sample before doing the next digestion to check that some of your nucleic acid is not RNA contamination from the bacterial host