restriction digestion - (May/24/2010 )

Hi, I am trying to get a ligation to work but in vain. Here is what I am doing:
One plasmid is pEGFP-N3 and the other is a spiroplasma plasmid
the two plasmids are cut with the restriction enzymes EcoRI and DraIII. I will be getting a 9027 bp fragment from one and a 1200bp fragment with the other. When I run the gel to check if the size is right, I am getting the right fragments. I am extracting the fragments from the gel and purifying them. The concentrations I get are about 12 ng/ul. Then I put in the two fragments in the ratios of 1:3, 1:1 and 3:1 and ligate them at 16C for 16hrs. I have been doing this from the past one and half month but can not get the ligation to work. One reason I am thinking is may be the concentration of the fragments. Because when I extract them from the gel, though I get the ng, there is no peak at 260/ 280 and the value at 260/230 is always around 0.3. I am not sure if this is the problem, but since this is the only thing I can think of, I was wondering if there is any thing that I can do to improve the quality of the DNA after gel extraction. If that is not the problem, can you suggest what I can do to to get my ligation to work.


Thanks in advance,
Sanu

Hi Sanu -- welcome to the BioForums!

I'm not familiar with the plasmids you're using, but are you trying to make a hybrid plasmid, or does one plasmid have an insert you're trying to re-clone?

If you're combining two plasmids, are they compatible with one another (e.g. each is from a different compatability group, such that they can co-exist in the same cell at the same time)? Or is your digestion such that you're leaving the replication origin of one of the plasmids behind? Is the selectable marker you're using remaining intact?

I assume you're concluding that your ligation is not working because you get no colonies after transformation. Are you sure your recipient cells are competent? Does the origin you're retaining work in E. coli (assuming you're transformng E. coli)?

As to improving the quality of your DNA after gel purification, you might try using a gel to which you have added 0.28 g/L (~1 mM) guanosine (e.g. Sigma G6752, FW 283.24) as a UV protectant to the 1× TAE used to cast the gel and as running buffer during electrophoresis. Stir with gentle heat until dissolved. This acts as a "sunblock", protecting against damage to your DNA by UV (see Grundemann, D., and E. Schomig. 1996. Protection of DNA during preparative agarose gel electrophoresis against damage induced by ultraviolet light. Biotechniques 21(5):898-903. There's a pdf here).

HomeBrew on May 24 2010, 03:28 PM said:

Thanks for responding to my question. What I am trying to do to remove GFP from pEGFPN3 and put it into the other plasmid pPOT and create a hybrid. I used the restriction enzymes which cut before the GFP gene and well after the gene. There are two selectable markers, one is GFP and the other is tetracycline resistance gene which is present in pPOT to which I am adding GFP. The pPOT plasmid has pUC replication origin, F1 and M13 replication origins. I have disrupted the F1 and M13 origins but the pUC origin is still intact. Also, I electroporated DH5 alpha cells. I also ran a gel to see if my ligation is successful today after I did not get any colonies on tet plates. I am uploading the gel for you. I am also confused about another thing. When I put my electroporated cells in LB+tet plates, I also put them on LB+tet liquid medium. I did all of them grew but none of them are flourescent. I ruled out the chance of contamination because the LB+tet liquid I used for the growth was still clean with out any growth.

Gel:
Lane 1 and lane 12: marker 10 kb ( lanes are 10, 8, 6, 5, 4, 3, 2.5, 2, 1.5, 1, 700, 500 and 300 bp)
Lanes 2-11: ligation product which is supposed to be 10,315bp.

I noticed that there are some bands around 2.5 bp and some bands above 10kb which I have no idea what they are.

Help me this. Thanks. Sanu

okay, as far as i get it you are trying to ligate the EGFP gene into spiroplasma plasmid backbone.

i would assume that the 2.5kb band is an indication for successful ligation of the product with itself (2x1.2=2.5kb) ...as can be seen in lanes 4 and 8-11. The upper band seems to be the chimera with the double insert (or triple insert?) ...it would be interesting to know the exact size of the band ...maybe you have an high range marker available that will give you that extra information?

To my point of view the ligation in lane 5 could show the correct ligation product since there is a minor size difference that could correspond to the 1200 bp. I don't know the difference between all the lanes (maybe you can provide us with that information) ...but it looks like that you increase the vector/insert ratio from the left to the right? Am i right?

To summerize ...i think that ligation is not your problem ...you have bands with double insert size and you have formation of bands that are larger than your vector backbone ...so everything should be fine

I would try to check the competency of your cells! What methode are you using? Chemically competent cells or electrocompetent cells? Selfmade or bought?

Transformation of large plasmids is not that easy (especially for chemical competent cells) ...have you tried to transform the uncut 9000 bp plasmid as a control? Does it work?
I think the problem is downstream of the ligation.

Good luck!

Regards,
p

pDNA on May 24 2010, 04:35 PM said:

Thanks for your input. I did not think about double inserts. I am using electrocompetent cells which I made. ( this is how I made them, as per the suggestion of a very intelligent lab member of mine who said her electroporation works all the time when the cells are prepared like this. I centrifuge 2ml of cells and resuspend the pellet in 200ul of cold distilled water. To this, I add varying concentrations of the ligation mixture and electroporate) I will try using electrocompetent cells from company and see if it will be successful. I am not very familiar with transformation and i had the idea that the electroporation works best. I did put in a control with the uncut plasmid but did not see any colonies. I thought may be the plasmid is not good enough because the plasmid that I extracted was about 10 days back. Any more suggestions are appreciated.

Thanks, Sanu

as you mentioned ...your positive control (uncut plasmid) resulted in no colonies ...this seems to be the core of the problem!!!

Concerning your protocol:
This seems not to be the best method for making good competent cells ...we in our lab do multiple wash steps with 1mM HEPES followed by a single wash step with 10%glycerol ...but protocols differ from lab to lab ...an actually very good online protocol can be found here.

Making good competent cells is "laborious" ...but when using a good protocol worth the effort!
I would trash your old protocol and stick to a new one!

Again good luck!
Regards,
p

I have used protocols in which ice-cold dH2O is the only reagent used to prepare electrocompetent cells -- but it was multiple washes (three or four, if memory serves). What time constant and actual voltage are you getting when you electroporate? Are the samples arcing?

In any event, I agree with the others -- it seems your problem is not in the ligation step, but in the electroporation step. You've got to get to the point where you get tons of transformants from your uncut plasmid control, or you'll have no hope of recovering transformants with your ligation mixture.

Thanks for your input. I will change the protocol for the preparation of competent cells and see if it works. Hopefully it will. Thanks a lot.