If there is no band after your digestion, the DNA is totaly degraded, or maybe there was not enough DNA to be visible on your gel. I Think you should take more plasmid DNA and fresh reagents for your digest.

good luck

Hi, I want know that the plasmid is original or was manipulated previously.

First be sure the plasmid is original and the recognition sites are not disrupted. sometime the MCS gets corrupted in cause of manipulation.

Second, you can increase the mix compounds concentration especially enzyme.

Third: Add 0.5 ul 100x BSA to each 50 ul reaction. most enzymes need BSA to doing function.

Forth: check you enzyme accuracy. Always dilute your enzyme in "enzyme diluent buffer" that is contains DDT or 2-mercaptomethanol and glycerol.

and Fifth; Check you gel and electrophoresis status.

Maybe a sequential digest is required. This is recommended for both enzymes taken from New England Biolabs and I agree with Babak, Bam HI needs BSA.

But now matter how you should see the (uncut) vector (I usually use 500 ng DNA which gives a nice signal), because you use the same plasmid (taken from the same tube) as for the control lane, right? If this is the case, your plasmid isolation should be fine, because uncut plasmid gives a signal and if there would be any inhibitory components, the restriction reaction would show uncut plasmid, too.

Maybe your digestion takes too much time? Both enzymes could cut elsewhere.

Alternatively you forgot to add DNA, or any component is contaminated with nucleases. I would renew all components.

Hi! Venkatesh

As per your discription your DNA sample has degraded may be because of star cativity ( altered specificity of restriction enzymes under non-standard reaction conditions) because you have mentioned the reaction mix used for the restriction digestion but not mentioned the incubation time. Even I have faced the problem of star activity while working with Bam HI.

The possible reasons of star activity are:

1. High glycerol concentration (> 5% v/v) which is not clear from your information as you have mentioned the volume of Digestion buffer but not the final strength used.

2. High concentration of enzyme/µg of DNA ratio

3. Non-optimal buffer in terms of pH, ionic strenghth, etc

4. Presence of organic solvents like, ethanol after DNA purification

5. Prolonged reaction time

So, please check the error in the earlier reaction mix and repeat the digestion keeping the above points in mind. Try to reduce incubation time.

Best of Luck

hi

1 unite enzyme is not enough to cut 1µg DNA. minimum I will use 5 unit, better even to go to 20 units ... 3 hours incubation should be OK

If DNA is absent following your digestion step, then you may have endonuclease contamination. Did you isolate your DNA from an EndA positive strain such as BL21? If so, this may be one major problem, although most DNA purification kits include optional steps to remove EndA. Recheck your propagation strain for its genotype and check your user manual for the DNA prep. Another possibility is that you are losing your DNA in a clean-up step after digestion. Does your protocol involve such a step or are you loading your enzymatic digest directly on your gel?

As mentioned by others, not all enzymes have compatible buffers to enable double digestions. For this enzyme pair (BamHI/HindIII), you will have better results with sequential digests. However, troubleshoot the single digests first (you should be able to see linearized vector on an agarose gel).

I have successfully used these enzymes to double digest pET28a in a 50ul volume- 1 ul each enzyme (results in the preferred %age of glycerol)~ however, my supplier of choice has been NEB. While I attempt to digest 1 ug in the reaction- I have NEVER been able to get the conc of MP DNA to be more that 60 ng/ul from a standard 5 ml culture miniprep so I am initially skeptical of your MP yields- assuming this is not the issue I have the following questions

-in your uncut lane- please clarify- do you see ONLY 1 band? you should see 3. what size are the bands running?

-when you perform single digests of each enzyme use the exact conditions accounting for the volume change with added water.

-you do not mention the temperature or time you allowed this reaction to proceed- I set this up end of day to cook overnight 37C

You have 1 ug DNA in 50 ul reaction volume. How much of that reaction are you running on the gel? You need at least 50-100 ng per lane to have a decent visualization of DNA on the gel. I suggest that you use at least 10 ug of plasmid in 50 ul reaction volume so that you have enough DNA to run on a gel and for further manipulation. The enzyme concentration of 1U/ul seems way too low. I suspect that you are diluting the enzyme. If you dilute the enzyme (in water??) you may lose some activity. Use 1-2 units per ug DNA. Another suggestion - when you run double digests, run controls alongside. Set up reactions of the same plasmid with individual enzymes. If it still does not show digestion, set up a control with plasmids that you know for sure contains the restriction sites to ensure that the enzymes are active.