Colonies grown without plasmid - (Oct/18/2013 )
Dear cloning experts,
I run out of ideas so I think it's time to ask help to the community.
I need to clone two chicken related transcription factors (TFs) into an rcas vector (avian retrovirus). I have first cloned these two TFs into an intermediate vector to add a tag on the 3'-end. I have checked my both constructs by sequencing, everything is fine.
I digest my constructs (1 kb) and my empty rcas vector (11.5 kb) by using 2 different restriction enzymes overnight, followed by a dephosphorylation for the rcas vector. I run my digestion products on a gel, cut the bands of interest and perform a gel extraction by using columns. Then, I ligate at 16°C overnight and transform competent TOP10 cells by heat shock. On the next day, I get from 5 to 20 colonies and I analyze them by PCR by using primers on the vector surrounding the insert location, in order to make difference between vector-only and vector+insert clones.
For one TF, I got my final construct after couple of tries but for the second TF, it keeps failing. I have tried many different conditions for the ligation (different ratios insert:vector
For this second TF, I get in general a few colonies and I don't amplify any product after the PCR with the same primer set. I use as positive control one of the positive colonies that I got for the first TF. I always get a band for this control, so it doesn't seem that the PCR fails. I have tried twice to do a few mini-preps from random colonies and perform a digestion test. I got an unique band for which the size doesn't correspond to anything that I should get...
The only reason that I have in mind would be that my cells catch the plasmid, get the resistance to the ampicillin and then release the plasmid. I don't see any other explanations (but I'm not a cloning specialist). I think that it's important to notify that I use competent-cell aliquots and LB+Amp plates from the same batches than my other coworkers, who don't have any trouble like mine for their cloning.
If anyone has any idea of what is going wrong, what could I try or test, or whatever, please share your ideas! I start losing my mind!! 
I will be very grateful for any help. Cheers,
Picui
Hi there,
i am not a cloning expert myself, but the only thing i can think of right now, is that perhaps, u overexpose your digested fragment to uv? Over exposure of UV can damage the sticky/blunt end of your band of interest.
i recommend u to start from digestion of your intermediate vector containing your inserts. From there, you cut bands of interest under 10sec.
I dont think is ligation or pcr fault, the other potential problem may come from transformation, have you checked the concentration of your fragments and plasmid?
in my experience, if you sure u have checked all possibilities and still u fail to get the result, its better to start all over again rather than u troubleshoot endlessly :)
I would definitely try this reaction without phosphatase treatment of your insert. You may see a slightly higher background, but over treatment can make ligation impossible.
Thanks for your comments.
To this concentration issue, I have try different amounts of insert/vector (molar ratio 5:1): 15/40 ng; 30/70 ng and 40/100 ng in 20-µL reaction. It worked much better when I combined 40 ng of insert and 100 ng of vector. So I then always proceed with these amounts for my next attempts/tests.
I can indeed try again without dephosphorylation. I did it once and got a lot of colonies, I screened 36 of them, all contained an empty plasmid. I should better screen more colonies this time.
Another way to reduce background without phosphatase treatment is to PCR very small amounts of your vector to produce linear DNA with cut sites of your choice at each end. Then, digest with DpnI and your enzymes to cut the original circular vector, and to produce correct overhangs on the PCR product. You need to make sure your primers have 5' bases past the restriction site to allow efficient cutting.
Picui on Tue Oct 22 11:48:42 2013 said:
Thanks for your comments.
To this concentration issue, I have try different amounts of insert/vector (molar ratio 5:1): 15/40 ng; 30/70 ng and 40/100 ng in 20-µL reaction. It worked much better when I combined 40 ng of insert and 100 ng of vector. So I then always proceed with these amounts for my next attempts/tests.
I can indeed try again without dephosphorylation. I did it once and got a lot of colonies, I screened 36 of them, all contained an empty plasmid. I should better screen more colonies this time.
What enzymes are you using?
phage434 on Tue Oct 22 13:18:27 2013 said:
Another way to reduce background without phosphatase treatment is to PCR very small amounts of your vector to produce linear DNA with cut sites of your choice at each end. Then, digest with DpnI and your enzymes to cut the original circular vector, and to produce correct overhangs on the PCR product. You need to make sure your primers have 5' bases past the restriction site to allow efficient cutting.
That doesn't make sense to me (assuming I've understood what you mean).
Amplification of the vector will generate linear DNA which is not phosphorylated. If this is digested with any restriction enzyme it will leave 5' phosphates (the two ends will self-ligate), thus dephoshorylation is still needed prior to ligating the PCR insert
The approach does work if you do the following;
(1) Amplify the entire plasmid (I usually do 25 cycles with Novagen's KOD proof reading polymerase - then gel extract) which will leave blunt ends which are not phosphorylated.
(2) Amplify your insert (you can include two blunt cutters in the For and Rev primers - e.g. SmaI, NruI, EcoRV), purify the product and digest with the blunt cutters to create a phosphorylated PCR product. Alternatively, treat a blunt PCR product with T4 kinase to phosphorylate your product.
(3) Ligate the two products in steps 1 and 2 (should be fairly efficient given the vector can't self ligate) and screen the clones. I like to screen by PCR (with oligos flanking the site of integration) as multiple fragments can ligate to one vector molecule. You can also include a few specific extra bases in one (or both) of the oligos used to amplify the vector. When ligation (of the correct orientation occurs) it will create a specific site which you can screen for.
*Amplification of the vector will generate linear DNA which is not phosphorylated.
* If this is digested with any restriction enzyme it will leave 5' phosphates (the two ends will self-ligate), thus dephoshorylation is still needed prior to ligating the PCR insert
Yes, but if the enzymes leave incompatible ends, you will not religate them. They will only ligate with compatible ends of your insert, which is what you want to have happen. You definitely don't need or want to do dephosphorylation. It will just cause problems.
Blunt end ligations are uncontrolled, and just are bad things waiting to happen.
phage434 on Fri Nov 1 21:09:00 2013 said:
*Amplification of the vector will generate linear DNA which is not phosphorylated.
* If this is digested with any restriction enzyme it will leave 5' phosphates (the two ends will self-ligate), thus dephoshorylation is still needed prior to ligating the PCR insert
Yes, but if the enzymes leave incompatible ends, you will not religate them. They will only ligate with compatible ends of your insert, which is what you want to have happen. You definitely don't need or want to do dephosphorylation. It will just cause problems.
Blunt end ligations are uncontrolled, and just are bad things waiting to happen.
Apologies i misread the original post. When I read the poster had been dephosphorylating the vector i'd assumed it was because the insert was going in on one site, so yeah I would agree that dephoshorylating is pointless.
Depending on the enzymes being used there could be self ligation of partially incompatible ends....guess it's not that common though.
I like the versatility of blunt ended ligations 