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!