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ds-oligo cloning/ligation issues - (Mar/06/2009 )

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Dear GilsonGirl:

I don't know if you solve your problem or not, but if you still with troubles, last year I have a very similar problem with an insert of 125 pb... the ds oligos that I use was from IDT to. Check this, maybe help:
a) Because the plasmid is to big is better that the ds oligos are phosphorilated and the plasmid dephosphorilated. That works for me.
B) because I forgot ask the ds-oligos phosporilated I liogate this into another plasmid (pGEMT), very easy a quick and then digest from this and ligate into my vector.
c) this is another kind of solution, for insert to short like yours, her we do PCR reaction with the insert in the tail of the primer, maybe you need 2 different PCR reaction for get that insert complete, but in time is more quickly and easy.

Good luck


Didn't want to start a new topic for this, so, here.

I'm going to move my insert from one vector to another, it's in Origene PrecisonShuttle system (using specific SgfI-MluI cuts on each side of insert) so it's should be a piece of cake, but I never really did classic cloning apart from TA cloning kits for PCR that 'just works'.

I'm waiting for the second vector now and planning it in advance. So this is what I intend to do:
Bought MluI and AsiSI (SgfI isoschizomer) from Fermentas (both FastDigest, they work in the same buffer).
I will make double digest of the first vector, short times are recommended as I read, so using 5-10 min at 37 degs of FastDigest enzymes should be enough. My vector is 8.6 kb from which 3.3 kb is my insert. I think around 300-400 ng should be enough for reaction.
I would run this on gel, separate the insert and isolate from gel with Qiagen MinElute kit (using as little of UV exposition as possible, changed to a safer channel). Quantify by spectrophotometry.

Now cut the new vector (7.1 kb) with the two enzymes same way. I don't want to purify such big fragment from gel, so I would dephosphorylate it only (there will be small 80 bp fragment between the two cutting sites, but as I understand it won't religate when dephosphorylated). Fermentas offers FastAP thermosensitive alkaline phosphatase that works directly in the FastDigest buffer, so I just add it there without purification, for 10 min 37 degs.
Then I inactivate the AP by heating on 75 5 minutes.

Mix the dephosphorylated second vector with insert in some molar ratios of (1:6 and mabe one or two others, count with the amount of second vector used for RE) and use Fermentas T4 Ligase (they state it should have 75-100% in the FastDigest buffer with 0.5mM ATP added, so I again add just enzyme and ATP) for 1 hour on RT. Inactivate by heating.
Transform chemically competent cells.

So this is the plan, we don't have AP or ligase, I have to buy them anyway, so why not use the compatible variants from the same company. I don't know if there are any traps in my plan.

What I don't know is what amount of the second vector to use for RE (but i would say like..100ng?), and what amount of ligation reaction to use for transformation, (up to 10% of cells volume?). Maybe I would make the second digestion reaction bigger volume than 20 ul, when I'm adding two enzymes, AP and ligase to it, so there wouldn't be so much glycerol percentage. I could probably purify after each step, QiaQuick colums can purify up to 10kb, but I don't know if that's necessary.

Thank you for any comments.


This sounds like an ok plan. I would make sure your RE digests were done in sufficiently high volume to allow you to dilute your DNA. substantially.

Ideally, you would find or make a vector with a different antibiotic resistance, so you could select against religation or uncut products from your insert plasmid. If you do this, you can avoid the gel isolation and purification. You can reduce background from uncut vector by using PCR to produce the (linear) fragment containing the vector backbone and whatever RE sites you need by adding them to the 5' ends of your primers. You must purify the PCR reaction prior to cutting (column). Then, you cut with your REs + DpnI to eliminate template plasmid (it will be cut since it has GATC sites methylated, that are cut by DpnI, whereas your PCR product has none). Heat kill the enzymes used for both PCR digestion and insert digestion. Mix the digested PCR product and the cut insert (with the cut vector, unpurified), ligate, transform.

You can prepare PCR product (uncut) if you doing a lot of work with a vector. This makes the sequence very quick and easy: Cut, heat kill, mix, ligate, transform.


These two vectors have different antibiotics resistance.
So I can cut first one and just add equivalent of 6:1 molar ratio to the second cutted dephosphed vector?
Wouldn't that lower the ligation efficiency or something?


Yes, it will lower efficiency, but so what? If your DNA and digestions are good, you will have plenty of transformants. The main difficulty is incomplete cutting of your vector and religation of the 80 bp vector insert. That's why using PCR for the vector prep works so much better at reducing background. Your real enemy is the colony which transforms, but has no insert. I'd gladly trade inefficient ligation and lower quantities for much lower background.


I see. The uncut vector and re-ligation is the main problem.

But I don't really like PCR much when talking about several kbs. If it was just ordinary bacterial plasmid, that selects out mutants in required genes needed for selection, then it may not matter. But my plasmid is a mammalian expression vector and I can't have it hit by some mutation in PCR, because it could then have lower expression that wild type for example, that would screw the subsequent experiments. The probabilty of this is low, but I would have difficulty to find it out.


Yes, that could be an issue. Use of high fidelity enzymes, especially Phusion, mitigates this, but if you really need to be certain, there is no substitute for sequencing your final version.


I see what you meant by the background. I didn't purify neither the insert nor the vector and made 3:1 and 5:1 ratios. Got pretty low colony count (around 30 on plate with 200 ul), only one of thirty tested colonies was positive for the insert, and that one had wrong size on the restriction. All others were empty.

I'm trying again on Sunday, gel purify both insert and vector and ligate in the ligation buffer this time, see if this approach is better.

Anyway I learned by this that I can have false positives on colony PCR from the remains of ligation reaction, so I guess it wasn't completely lost time.


Now, I digested old and new vector (Fast digest enzymes, 10 minutes), run them on gel, cut off insert (3,3kb) and linearised second vector (7,1kb) (used long wave UVA, tried to be quick as possible), isolated by Qiagen kit (MinElute for inserts and Qiaquick for vector, it's up to 10kb), measured concentration.
Made 5:1 molar ratio only, because the overal amount I got from gel was not enough (around 10 ng/ul), so I had 30ng of vector and 69 ng of insert.
Ligated 1 hour at 22 (Fermentas manual states 10 min is enough, but 1 hour can increase number, no PEG in buffer), and transformed XL1-Blue subcloning grade cells, that has been thawed once with 5ul of 20ul ligation reaction.
I got average 2 colonies/plate, most negative on colony PCR and the only possitive showed no insert on RE.
After that I remembered I forgot to heat kill the ligase so I repeated the transfromation next day with ligase inactivated, but numbers were similar, two PCR possitive clones has to be checked tommorow.

I'm trying to find the bottleneck. I will digest more DNA now, to have at least for 1:1 and 3:1 ratios using 50ng of vector, I will do single-cut-vector and single-cut-vector+ligase control. Also I used Fast AP to dephoshorylate vector right during the restriction, which was admissible in the manual, but now I will do it after digestion, heat kill.
But the main problem is getting low number of transformants. Bacteria are AFAIK fine, I'm using same for a month now, same transformation procedure. But I'm considering using those from non-thawed tube, that should increase the efficiency. On the other hand there are only two left (handy 500ul packing ) and I have three inserts (same size, different mutation) and the difference shouldn't be such high.

Is there anythyng else I can do different? I can hardly do something with the UV step, but I think 400 - 315 nm shouldn't cause considerable problems, or would the potential damage be compensated by higher amount of DNA in ligation reaction, like 100 ng of vector and appropriate amounts of insert? Ligase and enzymes are all new so they should work, restriction seems fine judging by the bands of first vector with insert. The concentration ig gel excised fragments could be checked on gel, however this time the concentration was too low to be seen, for same reason there is probably no point in putting the rest of ligation on gel.

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