Problem with separating linearized/circular plasmids for cloning (or problem with CIP maybe)?

I'm doing molecular cloning at the moment. I need to insert a 3300bp fragment into an 9000bp vector (carrying an Amp resistance) via a BamHI site. The insert does not carry a resistance of course. The problem is, that I get essentially the same amount of colonies on my agar plates for ligation as well as for the controls. The vector has been linearized (by BamHI), CIPPED (= treated with calf intestine phosphatase, CIP), put on an agarose gel to separate linearized from not linearized molecules and then extracted again.

[more detailed: I digested 2µg of vector with 0.5µL BamHI-HF in a 20µL reaction volume for 2 hours. Afterwards I added 0.5µL CIP for 1 hour, which should be more than enough.]

For transformation (in E. coli DH5alpha), I used the following samples:

ligation: vector + insert + ligase
control 1: vector + ligase
control 2: only vector

As I said, the negative controls show roughly the same amount of colonies as the ligation and control plates. To me, that implies that either there was an insufficient separation of linearized vs. circular vector or that CIP treatment did not work. However, the same tube of CIP has been used for other purposes where it worked. On the other hand, I cannot quite imagine that there should be a significant amount of uncut vector in my extracted sample, because I did a pretty extensive separation and tried to cut really precise and tiny cuts.

Any suggestions what I could try or change in my procedure?


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  • Estanis Navarro · IDIBELL Bellvitge Biomedical Research Institute
    Johannes, comparing the number of colonies is not enough to say that you have failed in your cloning. Test several colonies by colony-PCR, perhaps you were lucky enough, you've got clones, but you don't know it.
    If you really failed to clone your insert, try Shrimp Alkalyne Phosphatase. It is thermostable and after treating your plasmid you heat-kill SAP and can continue with your cloning.
    On the other hand, trying to clone a 3,3 kb insert in a 9 kb vector is not trivial!!! I would also use the highest competent cells available (there are commercial cc that yield up to 10e8 cfu/ug), or transform by electroporation.
    Have a good cloning!!
  • Daniel Cohen · University of Pennsylvania
    Johannes- two scenarios come to mind. Based on how many colonies you have in your negative control, you can discriminate between these:

    1) Lots of colonies in your negative control. Implies that either your CIP treatment didn't work or you have uncut plasmid in your prep. Why? Restriction enzymes sometimes hang on to DNA even after cleavage, which might sterically interfere with CIP. You might have better success if you remove BamHI before CIP treatment. Second, and more likely, your supercoiled DNA did not resolve completely from your linear DNA in your agarose gel. Use a low % agarose (0.6-0.7%) and run for at least 2 hours before gel extraction.

    2) Very few transformants in either plate. This implies your digest and CIP treatment is OK, but your ligation with insert is bad. Possible causes are damage to your vector end by excessive CIP treatment (chewing off of your sticky ends), damage to your vector by excessive UV exposure, incomplete digestion of your insert with BamHI, or poor DNA quality of your insert preparation. Most frequent culprits are excessive CIP or UV, both of which are easy to fix (use NO MORE than 0.5 units of CIP per microgram of DNA).

    Good luck!
  • Johannes D. · Johannes Gutenberg-Universität Mainz
    thanks for your suggestions.


    correct me if I'm wrong, but I guess since I have roughly the same amount of colonies on ALL the plates, I would most likely have to pick a very large number of colonies to screen with a PCR or a digestion to even obtain ONE correct one...


    I have really many colonies on all my plates (about the same amount on each), so ligation should be fine - or maybe ligation is not working at all PLUS vector linearizing ist not working at all, too, so all I see are colonies with unligated, unlinearized vector...but that's not too likely, I guess (?)

    I separated the linearized/circular vector on a 1% gel, applying 125V for about 4 maybe I should use lower percentage (0,7%) - on the other hand, I had a pretty reasonable separation (on the printout, I can basically see a very weak band at around 11.000bp or something plus a very strong band at the exact size where the linearized form should be.
  • Daniel Cohen · University of Pennsylvania
    So it might be the case that your CIP treatment was not adequate. You should try a transformation control with vector +/- CIP treatment to check.

    To improve separation on your gel, don't overload the lanes. Tape several lanes together so that your band (single cut vector) is long and narrow rather than short and fat. No more that 1ug per lane of an 8-lane gel.

    Even if your cut vs. uncut vector look like they are resolved, the DNA runs as a distribution, and your "band" is the peak intensity, and does not show where the tailing ends of the distribution lie. To improve separation, I'd suggest using the lower %age agarose and run at a lower voltage (65V).

    Finally, your cloning will be greatly simplified if you using directional cloning with 2 different restriction enzymes. Although this may seem like more work, it obviates the need for CIP treatment and may save time in the long run.
  • Estanis Navarro · IDIBELL Bellvitge Biomedical Research Institute
    Johannes, yes, you should pick a relatively large number of colonies.....
    On the other hand, the fact that you have many colonies in your vector (no insert) control means that either you have a carry-over of non-digested, supercoiled vector (that transforms much better than ligated plasmid) or that your CIAP treatment didn'work and you religated your plasmid (intramolecular ligations are more efficient than intermolecular ones).
    What I would do:
    -Perform Bam digestion in 50 ul final volume with 1 ug plasmid, 5-10 units of enzime and digest for 2-3 hours.
    - Treat the digestion with SAP. I prefer using SAP over CIAP because SAP can be readily killed by heating and you don't have to do lots of phenol extractions or column/gel purification to get rid of CIAP.
    -ligate overnight, at 14-16 C and in 10 ul final volume, and transform. You should expect no colonies in your control if Bam digestion and SAP treatment worked well.
    Finally I agree with Daniel's advice of using directional cloning. This is always the best approach.
  • Joan Lemire · Tufts University
    Could it be that the BamHI itself is blocking the phosphatase? It does not release easily. This is from the New England Biolabs site, about their "BamHI-HF"
    "The increased specificity for the BamHI-HF™ cut site has increased binding of the enzyme to the DNA and the enzyme may remain attached to the DNA during gel electrophoresis. To disrupt binding, add SDS to a final concentration of 0.1% - 0.5% or purify DNA before electrophoresis."
    I would clean up the digest with whatever spin column you use to clean up PCR , then resuspend in restriction enzyme buffer for the phosphatase reaction.
  • Johannes D. · Johannes Gutenberg-Universität Mainz

    hmm, good point - but wouldn't heat inactivation of BamHI (65°C for 20min or something) do the trick as well? Because I'm already in the middle of trying that...
  • Praveen Singh · Universidad Autónoma de Madrid
    I have done many cloing by single enzyme cutting and without treating vector with phosphatase. I found sometimes treating vector with phosphatase can harm your vector and you won´t get any or very few transformants. Why don´t you try cloning directly withput treating your vector with phosphatase and use more amount of vector and insert ratios. Try 1:20 ratio and directly do colony pcr. Do not waste time on discussing about cloing because cloing is many time based on your luck too..
  • Daniel Cohen · University of Pennsylvania
    BamHI is not heat inactivatable!!! You have to use a digestion clean-up kit (QiaEX resin or Qiaquick column or equivalent product).

    In contrast with Praveen's experience, I have never seen successful cloning of an insert into single cut vector without CIP treatment. The self-ligation frequency is simply too high. Maybe he was very lucky to be working with extremely small inserts where a 20:1 ratio is easy to achieve. It will never happen with a 3.3kb insert, and would be a very risky strategy in my opinion.
  • Johannes D. · Johannes Gutenberg-Universität Mainz
    @ Daniel: damn it, I was mistaken when I looked on the poster with the enzymes in our lab, stupid mistake - BamHI is indeed not inactivated by heat. So I will try the SDS treatment prior to CIP treatment...

    @ Praveen: the problem is not that I get few colonies but rather that I get many colonies (but not significantly less in the negative controls) CIP damaging the vector should not be the problem...
  • Daniel Cohen · University of Pennsylvania
    @Johnannes- Mistakes like that happen to the best of us. If you can't get the CIP treatment to work, switch over to Gibson cloning. Since it does not involve a ligation step, you don't need to worry about self-ligation. Moreover, Gibson cloning has the advantage of controlling orientation of your insert, which the restiction enzyme-based cloning does not in the case of a single enzyme digest.
  • Ria Goswami · Southern Illinois University School of Medicine
    I think two possible explanations could be....your CIP did not work or you have a little amount of uncut vector there.But since you say the CIP works fine..I would guess the second one is the case.
    You can try two options:
    1) either try to cut the vector and run it long enough comparing with the uncut vector and try to seperate cut from uncut
    2) Try to do PCR screening with one primer inside the insert and one outside.
    I think it will solve your problem.
  • Christopher Provost · New England Biolabs
    You could also try Antarctic Phosphatase from NEB. I have pretty much stopped using CIP with occasional exceptions. 1 ul of Antarctic Phosphatase will dephosphorylate up to 5 ug of vector DNA and it can be heat inactivated at 65ºC in 5 minutes.
  • Conrad Quinn · Centers for Disease Control and Prevention
    BamHI overhangs will self ligate and transform DH5-alpha cells quite efficiently without an insert and even with CIP treatment. I wonder have you screened the transformants for inserts? Irrespective of the number of colonies you're getting on the control and test plates, you may find that the cloning has indeed worked when you screen. Alternative strategies would include increasing the amount of insert in your ligation mix or using dissimilar ends for your cloning.
  • Nigel McLeish · University of Manitoba
    I think if you have to digest for 2 -3 hours with a HF enzyme, then the enzyme isn't working properly. I would get new BamHI.
  • Pablo Becker · King's College London
    Are you Amp plates fresh? I am almost sure you check this, but I am just asking because sometimes the problem is the obvious one. Did you check with DH5a without plasmid to be sure they do not grow?
  • Matthew Moreno · Vical
    hi, some people do not use Ampacillin due to satellite colonies. Make sure you k now how to tell satellite colonies from others. You can also increase the Amp (100 ug/ml) in the media or switch to a Amp analog like carbenicillin.
  • Donna Williams · Johns Hopkins University
    I find that I have a much better signal to noise ratio if the gel purify my vector twice.
  • Adrienne Smyth · Worcester State University
    What I would do is cut a much smaller amount of plasmid, say 200ng , forget separating it on gel, phenol extract, EtOH ppt, and then ligate your fragment in- try 600 ng to get approx = ends and use 1/10th the amount in your transformation. Pick 12 colonies, plasmid prep, and run uncut on get - slower one should have insert. Run uncut plasmid of approx 9 kb alongside as marker, and the original cloning vector of course) Pick selected positives and confirm by cutting with BamH1 to ensure fragment there. Best of luck
  • Fedora Sutton · South Dakota State University
    Do you know the sequence of your insert? If you do, is there a second enzyme that will work so that you can perform directional cloning and at the same time not have to bother with CIP treatment.
    Alternatively before you make any other changes, it is worth addressing Pablo's and Matthew's comments about your amp plates. Regarding the use of ampicillin, we switched to carbenicillin. The bla gene also confers resistance to carbenicillin. However carbenicillin is more stable and does not allow growth of satellite colonies.
    Just in case your Amp concentration is too low and you are viewing satellite colonies as transformants, I agree with Matthew that it would be worth switching to carbenicillin.
    all the best
  • Patrick Van Gelder · Ghent University
    I would , like already suggested switch CIP to sea-born phosphatase that can be heat-inactivated (I used to work with shrimp alkaline phosphatase from Boehringer), may be your CIP is dephosphorylating also your insert (it is hard to separate it from the plasmid prep.
  • Koen Venken · Baylor College of Medicine
    Although I use a negative control, I generally still check colonies when the negative control gives an equal amount of colonies. Sometimes you even get much more colonies on the negative control compared to the true ligation. That is a good indicator too, since that suggests competition of the true ligation with the self ligation. In the latter case, most of the colonies on these ligations contain the desired insert. So, a negative control is informative, but not the golden rule. You should still analyze some of the colonies.
  • Patrick Van Gelder · Ghent University
    Koen is right, had several times the same experience
  • Vik Rampersad · Samuel Lunenfeld Research Institute
    Since you say you get the same number of colonies on both negative controls as your ligated transformation, this suggests that teh BamHI digestion is incomplete. When you run DNA on an Agarose gel there is always some "tailback" of the faster running product so you could have some uncut vector in what should be a linearised band. What % Agarose did you use to separate the bands and more importantly, how many bands did you see? 2 or 3? If 3 then the enzyme is definitely not working well at all as you are seeing uncut (supercoiled) at the bottom band, linear at the middle and nicked at the top.
    I would repeat the digest using new enzyme and maybe switch to Shrimp Alkaline Phosphatase (heat inactivatable)
    I would also still screen your ligation colonies as the insert may well be in there amongst the background, but its probably better to simply start from scratch
  • Thomas Macartney · University of Dundee
    I would change the protocol and include an additional restriction site other than BamHI to allow directional insertion of your fragment. As for your current problem it does look like your BamHI is not cutting efficiently or the phosphatase is dead, there should be no need to have to extract twice on a gel. You should definitely be obtaining no colonies on the negative control.
    I would recommend fresh BamHI and shrimp alkaline phosphatase. Perform the digestion with 1.5 ug vector, 2ul BamHI, 2ul SAP (shrimp alk phos), 10ul buffer and dH2O to 100ul. Cut for 2-3 hours at 37oC, it is fine to add the phosphatase at the same time as the enzyme. Run the whole lot on a couple of adjoining lanes and extract the linear band, eluting in 50ul. Use 1ul per ligation and you should be fine.
  • Ronald Godiska · Lucigen Corporation
    You mentioned that you added CIP, but not CIP buffer. This buffer is usually high pH, far different from restriction buffer. After the restriction digest is done, add 2 ul of 10X CIP buffer plus 0.5 ul CIP directly to the reaction, incubate 60 minutes at 37C, and purify the vector (e.g., on a Qiagen column).
  • Dasheng Zheng · Wuhan Institute Of Virology
    Hi, according to my experimental experience, the BamHI is so active that it will over-cut if incubating over 30min. So, please try: excess BamHI in the restriction system but only incubate no more than 30min, and inactivate BamHI by heating or EDTA, purify the reactional products and then dephosphate/ligate.
  • Robert Marano · University of Western Australia
    Most people seem to be concentrating on the digestion, CiP process, ligation etc. However, I think the opinions by Pablo Becker and Matthew Moreno are probably the best and easiest to discount. If there are the same number of colonies on each plate then it would seem that antibiotic selection is not functioning. Were the plates made recently, was the Amp relatively fresh or had it undergone a number of freeze thaw cycles etc. When adding the Amp do you add to the melted agar or spread on top. If added, and the agar is too hot it will deactivate the Amp. As suggested earlier, try plating just the DH5 alpha with no plasmid

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