I have a vector with the restriction sites PAC1/SGS1 that I plan to use in order to ligate my insert which does not have these sites on it.
In order to make the sites I used two primers (one forward, one reverse) at 10uM concentration that contain the beginning and end of the sequence of my insert with the PAC1/SGS1 restriction sites added on the ends. A few nucleotides were added past the restriction sites in each sample which was suggested by my professor who got results using this procedure with the same sequence. However he was not placing the PAC1/SGS1 restrictions sites specifically.
The PCR samples, for the insert, have a total volume of 25uL and used the following program setup:
I have confirmed through electro-gel phoresis & gel purification that I retrieve ample PCR samples at the correct band size. Agarose Gel was 1% in 1X TAE solution and ran 100V constant for 45min.
Both the vector and insert, with new restriction sites form PCR, were then digested using PAC1/SGS1 in a 20uL volume and incubated at 37C for 20min using a warm water bath. Reaction mix is as detailed below:
- 1uL of enzyme (one with both enzymes, another with 1 enzyme)
- 2uL of Green Buffer
- 1ug of DNA
- ddH2O to fill to volume requirements
I confirmed using the same gel procedure as above that the desired bands had been obtained and extracted/purified the samples from the gel for use in the ligation.
The ligation uses T4 Ligase and had the following reagents:
50ng Vector DNA
- 1:1, 1:3, 1:5, 1:7.5, & 1:10 Insert DNA (Respective to Vector amount)
- 2uL 10X T4 DNA Ligase Buffer
- 1 Weiss U(5 Weiss U/uL stock) T4 DNA Ligase
- ddH2O to fill to total volume if needed
I observed colonies in my positive control and no colonies in my negative control. I also observed no colonies in my other samples except for a single colony in the 1:3 and two colonies in the 1:5. I retrieved some samples of the colonies for a plasmid prep:
I retrieved the overnight inoculations and used a plasmid prep kit in order to retrieve DNA sample and measured the samples on a nano-drop to determine the purity. The absorption rates came out fine.
The samples were then digested with PAC1/SGS1 using the same procedure as above.
The digest samples where then run through a 1% Agarose Gel in 1XTAE solution set to 100V constant for 45min. A sample of digested vector/insert were also run in the gel in order to compare band size to the ligated samples.
The gels have always resulted in a recombination that seems to be a common way for the XL-10 cells to form resistance to AMP and proliferate. So far I have not seen ligation of my insert/vector.
I sent the insert sample to be sequenced however the primers were not able to run far along enough to determine if the primers placed the proper restriction sites on the insert.
I am currently going to try another ligation which I will then digest again to determine if the ligation is working properly. I am also creating new competent cells just in-case the ones I am working with are not able to take up the plasmid due to loss in competency. I have also ordered new primers with no extra nucleotides at the end to see if that may also be an issue.
I have a feeling that the problem may be the extra nucleotides although I don't know why it would've worked for my professor who carried out a successful ligation before me doing the same thing. The plasmid I am using to retrieve the insert is actually from his ligation.
Might there be something I am missing here? Could it really be as simple as removing the extra nucleotides at the end of the primers or is there a problem with my methods?
Francesco, PacI seems to cut well even with only a 1-nt extra overhang but I haven't been able to find any data on how well SgsI(?) cuts near the ends of linear DNA. Did you check with the manufacturer to see how well it cuts near the ends, and how many extra nucleotides did you include following your restriction site?
It sounds like your enzymes are working since there is no evidence of significant recircularization. of your vector from the transformation results. How large is the insert?
Hi there,
First, do you purify PCR product before digest? Digest may be done on raw PCR product but the efficiency is less than on purified product.
Second, you are mentioning Green Buffer for digest so I guess you are using ex-Fermentas enzymes (now ThermoFisher/Life Technologies)... But none of the two enzymes is actually fully active in the Green Buffer according to this webpage:
https://www.lifetechnologies.com/fr/fr/home/brands/thermo-scientific/molecular-biology/thermo-scientific-restriction-modifying-enzymes/restriction-enzymes-thermo-scientific/conventional-restriction-enzymes-thermo-scientific/reaction-conditions-for-restriction-enzymes.html
PacI is only fully active in its specific buffer and Sgs1 is fully active in the Tango Buffer... So I would suggest a sequential digest for optimal efficiency.
Hi Francesco
After the ligation step you can check the presence of insert by PCR. Screen the colonies by PCR first, amplification would confirm the presence of insert. You could use a primer that goes across the PAC1/SGS1 site and another primer in the vector. After this step a plasmid prep and sequencing can verify the successful ligation.
To question number 1 from mark: My insert is around 1,100kb and I haven't seen any evidence to suggest it recirculized. I should probably ask the manufacturer about SGS1. SGS1 also used to be called ASC1 I believe so that might be why there isn't info under the new name.
To number 2 from Dominique: I have done the digests in both Green Buffer and regular FD buffer from Thermoscientific but I haven't heard of Tango Buffer before I will check on that. I have done a PCR purification before digest as well (didn't mention it here) and the results were the same. I've done this with a variation at each step just to make sure. I even created entirely new reagents for the PCR just in-case something there was wrong.
To question number 3 from Indu: That's a good idea I'll use that to check the presence, I was also thinking of just running a digest on the ligation product and running through a gel to see if anything did ligate.
I'll update after trying these suggestions, thank you very much.
What I am trying now is some different primers with poly A tails as well as 8-12nt attached beyond the restriction site on the 5' end. I made sure to leave at least 15nt on the 3' end to attach to the target sequence.
Using this site https://www.embl.de/pepcore/pepcore_services/cloning/pcr_strategy/primer_design/ I was able to determine this is the best course of action and so the truncated primers I created with no extra nt on the 5' end are not useful to the experiment.
Also I am using G. FD Buffer which according to the thermoscientific website is 100% efficiency, sorry for that confusion.
I will be checking the ligation step soon now that I have the new primers, thank you for all the help.
The PCR with the new primers showed some very light bands above mCherry suggesting that there is some sort of ligated plasmid there.
I am checking the ligase itself right now to see if I should bump up the amount used and afterwards will be performing another ligation, with the new primers for PCR. The only difference this time will be the amount of PCR product I create as in previous runs the concentration has been very low (less than 20ng/uL) and is probably the reason why the bands are so faint.
Francesco, thanks for the updates. I've had good success subcloning with NEB's AscI and PacI in the past.
No problem Dr. Chee. I will be updating this until I obtain the goal product.
The ligase experiments showed no reason to think I should increase the amount used. However when comparing with a phosphotase treatment, the ligations using the mCherry Vector that had the 5' phosphate removed seemed to produce stronger bands.
Currently I am creating more concentrated samples to retrieve more product from the ligation, since the amount retrieved is 10x lower compared to plasmids that have not been run through a ligation reaction. Once that is done I will be comparing two different ligations, one where sample vectors have intact phosphates and one where they are exposed to phosphotases.
Francesco, I'd like to find out how much vector DNA you add to one of your ligation reactions, and how much of your ligation you use to transform your competent cells with.
Usually I add anywhere between 20-100ng of Vector with Insert concentrations 1X/3X/5X/7X/10X higher than the vector. After that I use 5ul of my ligation with 50ul of XL10 Gold cells and transform the entire 55ul reaction mix onto Agar LB plates.
I usually run the ligation for 10 min at room temperature but today I will be running differing times to see if it helps as well as with a phosphotase reaction on some of the vectors.
A good note is that for most of these reactions my insert concentration samples were incredibly low so there has never been a lot of ddH2O in my reactions because there was virtually no room for it. The latest ligation that showed some bands I had a better insert sample so I'm trying to see if running the ligation longer will create a stronger band.
Update for now:
I have been using the pGEMT-easy vector from Promega. This vector acts similarly to blue-script in which colonies can easily be screened (blue meaning no insert, white meaning insert). My plates all contained white colonies (I thought 100% efficiency was a bit strange but perhaps that is normal) and have begun testing them to see if the proper insert is there. Initial gel phoresis is showing promising results with the STN1 1.1kb insert ligated into the pGEMT-easy vector. I will be sequencing soon with T7 and SP6 primers.
I decided to use this system I have been told by various cloning experts that using just a PCR product with restriction sites on the end is a more difficult method to get working so I decided to find a cloning vector.
The next step is to clone in the insert into the pcDNA 3.1 mCherry vector and test using phoresis and sequencing. Afterwards the desired product is more or less achieved and ready for use.
After checking the cell colonies through restriction digests with the enzymes Pac1/Sgs1 the results show me that the restriction sites are not there. I have also gotten bands similar to previous ligation that ended up being recombinants that the XL10 gold cells used to get around the ampicillin.
I am currently sequencing the plasmid from the colonies and running yet another digest to check if the pGEM-Teasy vector is present or if the old restriction sites for STN1 were kept and the new ones were for some reason not place on there by PCR.
Will update with results soon.
So the pGEM-Teasy system did not work either. Somehow the insert was placed in the vector but it did not have the restriction sites we wanted, instead it kept the old restriction sites. Apparently the primers may have been mixed up in such a way that a PCR product with the old primers was used instead of the ones with the proper sites for digestion.
The major problem seems to be that the PCR products themselves have very low concentration even when the signal from UV imaging in the electrophoresis gel is very high. We suspect there may be a problem with our nanodrop machine and will be analyzing the gel samples a different way.
Hopefully that fixes the problem. Thank you all for the help.
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