Hi Wubliker,

It sounds like your transformation may not be optimal. Are you doing heat shock for transformation? Also when you got your colonies, were these colonies obtained following selecton? 

Hello Augusta,

Yes I did heat shock and used ampicillin as a selection marker and it proved to me that  in the control plat there was no colony.  

Because the empty pUC18 itself has Amp marker, it's possible that some of the colonies do not have insert. To check this, when you extracted the plasmid DNA from the colony, you can run it on agarose gel and also include a lane with pUC18 alone. The DNA that contains insert should have a higher MW than the empty pUC18. 

There is a possibility that your insert is there and perhaps lost the restriction site. If the extracted plasmid DNA has the correct MW but apparently no product when digesting with XbaI and EcoRI, you can perhaps try another restriction enzyme, preferably one has a cut site within your insert and another cut site within the vector.

Yeah, you are right. But I suspect the problem might be on the ligation reaction.

That's good trouble shooting Jamin.

Hi Wubliker,

First, XbaI digestion is blocked by dam methylation at GATCTAGA and TCTAGATC sequence, so check the sequence around XbaI in your Actinobacillus fragment (If one of these sequences is present, it will be cut in a PCR fragment but not on the chromosome or on a plasmid).

Second, since you say you get lots of colonies, I think the problem is not transformation (unless you reuse electroporation cuvettes that have not been properly sterilized, or you use old antibiotic plates in which the amp concentration is too low )

About the ligation, 2 possibilities:

1. The DNA you get in your numerous clones has very different lengths and digestion profiles: even if the two EcoRI and XbaI extremities are incompatible, you can get colonies from your cut vector alone (stemming from recombination events or ligation of extremities nibbled by contaminating nucleases....). But the amount of these products should not be huge compared to your expected product if the ligation is performed in the right conditions (i.e. vector/insert 1:3 in moles).

2. The DNA you prepare from your transformations looks mostly like the starting vector: I would bet you have uncut vector. If both XbaI and EcoRI are uncut the plasmid will be supercoiled and transform way better than your ligation product. Even if just one of the sites is uncut, it will need just one intramolecular event to religate whereas the desired product requires first an encounter between two molecules (vector + fragment) and then the same intramolecular event.

So check your restriction enzymes on fragments giving visible size changes upon digestion. For instance if you have already a pUC plasmid with a fragment inserted, use it as a vector rather than the pUC itself: cut with the two enzymes (one at a time if you want to be sure), purify from an agarose gel the band corresponding to the vector, and use this as your vector source.

Hope it helps

Olivier

Hi Wubliker,

Ages since I've done any cloning, but I always liked to dephosphorylate my vector so you get fewer vector-only colonies on transformation. Gel purification gets rid of any uncut vector too.

Also a quick look at pUC18 map shows the LacZalpha part present, which means you should be able to do blue/white selection with XGal.

Can you do a vector only ligation control to see if you have high background?

best

Bryce

Thank you so much all of you for your concern and dedication. Its really helpful and makes me to learn more and more from you scholars. I will use combining all your suggestions.

@Olivier, after I have extracted the plasmid from the colonies, I did 3 digestions (1 with both REs and the other 2  with XbaI and EcoRI). And in all 3 conditions, the result was all the same. So, I think, as far as the RE is concerned, it could digest perfectly.

Hi Wubliker,

And what is the result? I guess it's just one band, corresponding to the linearized vector . When you digest with both enzymes, the EcoRI and XbaI sites are so close that you can't see the difference in migration.

So, if the colonies you are talking about are the transformants, and if you only get the starting vector in these transformants, there is still the possibility that you have just 1% uncut vector and even if you can't see it on a gel, it will appear in the transformation because of the differences in ligation and transformation efficiency, so you could try to increase the digestion time for instance.

Otherwise, there must be a problem with your insert:

either its concentration is overestimated, or the XbaI site is not cut because of dam methylation or it produces a toxic protein, or you put too much EcoRI and got star activity. Did the fragment look OK on a gel?

There is one last thing I just thought about: did you inactivate your enzymes (by heating) before the ligation?

Olivier

You may also want to consider the size of the vector that you are adding. You may want to increase the concentration of digested vector used to increase the chanced that your relatively large insert will actually insert into the vector. Also make sure that there are no restriction sites associated with your restriction enzymes that would result in the loss of the ampicillin resistance gene. After you ligate the insert into the vector run a test digest on an agarose gel next to digested vector or undigested of both to see if insert has been added before transfection. Hope that helps.

@Olivier, yes, in all 3 conditions, I have got single bands with the same MW. As you said it,these two RE sites are close. And now I will try with longer digestion time. And, no,  I did not inactivate enzymes by heating before the ligation.

@Alec, now I am doing it with higher vector concentration and longer ligation time. We will see whats gonna happen.

Really thank you both!

Hi Wubliker,

you should absolutely inactivate your enzymes one way or the other, either by purifying your fragment and your vector on an agarose gel or by heating (65°C 20 min) the restriction reactions, otherwise, the enzymes will redigest your ligated products during the ligation. If you heat your digests, don't put too much digest in your ligation because salt inhibits the ligase, if you purify on a gel, you don't have to bother wit that.

I don't agree with Alec: you must increase your chances of bimolecular ligated product  over the uncut or religated vector, so don't increase the vector/insert ratio. As I said before, the ratio generally used is 1 vector/ 3 insert in molecules, which given the MW you provide gives a 4.2 weight ratio in favor of the insert. If you purify  your fragments on a gel you could even increase that ratio, but first you should get rid of all these colonies containing your vector alone so digest more and do the vector only ligation control recommended by Bryce. And XbaI and EcoRI are in the polylinker of pUC18, so they don't cut anywhere else in the plasmid.

Olivier

Hi Olivier,

Here is how I did it.

Vector

plasmid---Digest with XbaI and EcoRI-----Gel extract or PCR clean-up

Fragment

PCR----Gel extract----Digest with XbaI and EcoRI-----Gel extract or PCR clean-up

OK, then the enzymes are not there anymore in the ligation. So, do extensive digestion followed by the gel extract procedure in order to be able to increase the insert/vector ratio. Use fresh ligase and fresh ligase buffer (contains DTT which decays over time). If it does not work that way, it might be that your fragment is toxic. Then try a lower copy number vector.

Olivier

Hi, Wubliker.

If blue-white selection and others mentioned above tips won't work, it may be useful to change cloning strategy. For example, cloning fragment may be divided into set of smaller fragments, that will be subcloned separately, one by one.

Igor.