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Electroporation doesn't like my plasmid???


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#1 Mattise

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Posted 11 August 2010 - 01:53 PM

Hi all, new user here. Looks like a very helpful message board!

I am trying to clone a plasmid using electroporation and am having some difficulties. The vector and insert are both ~2.2kb, the vector being a derivative of pUC18 minus the alpha fragment of the LacZ enzyme. The insert is for a beta-gal gene.

Here is my current protocol.

Vector is taken from a plasmid prep, insert is taken from a gel purification of my PCR product.

Vector and Insert both digested with PstI for 1hr. at 37C.
Shrimp alkaline phosphatase added to the vector, and both allowed to continue incubating for 30 minutes.
Enzymes are heat killed at 65C for 15 minutes.
A T4 ligase mixture is made from these and allowed to go overnight at 16C.

At this point, if I use Z-Compenent cells that I have made, I can transform with decent efficiency. I get around 50 blue clones per plate this way.

However, when I do electroporation, I get zero clones. I electroporate both pUC18 and my vector (pre-digest of course) and both give me lawns of clones.

I have tried gel purifying the ligation reaction but still have the same issue.

Kinda stumped on this one.

I don't know my DNA concentrations, but I do know that it works with the Z-competent cells, so I wouldn't think anything needed to be changed there.

#2 perneseblue

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Posted 11 August 2010 - 08:01 PM

Please correct me if I am wrong.
I understand that this is a simple ligation between a vector and insert where both DNA fragments have been cut with PstI.

The first thing to do is to check your ligase, T4 DNA ligase and its buffer can easily go off. The ligase buffer has a strong smell (DTT). If you can not smell anything from the buffer it means that the DTT has been oxidized and the dATP is probably oxidized too. Bin the buffer and get a new one. T4 DNA ligase does denature at -20C.

To check that the ligase is working, do the ligation reaction between your insert and vector. Run part of the ligased DNA mix onto a gel. You should see several high molecular weight bands.. which are your ligated DNA products. If you do not see those high molecular weight bands, the ligase is dead. Bin it and buy a new one (Buy the small vial. In my experience, a lab can rarely finish a small vial before it becomes unusable.).

If you do not see, any DNA, it means that the DNA was lost during the ethanol precipitation or the 70% ethanol wash. I find that if you centrifuge the sample after adding the 70% for 12 min, you can pellet down any DNA that got detached from the tube well when the 70% ethanol was added. I also find that doing the ethanol precipitation in a 500ul tube is very helpful. A strong light can be used to help see the DNA pellet.

Oh, and did you check to see if the cut vector and DNA fragment are actually present? Perhaps by gel or nanodrop? It is possible that the reason why there are no colonies, is due to the absence of DNA, which may have been lost during the purification step.

What was the ratio (mol) that was used for the ligation?
May your PCR products be long, your protocols short and your boss on holiday

#3 Mattise

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Posted 12 August 2010 - 08:45 AM

Thanks for the reply. I know for a fact that I have ligated plasmid because I used the exact same ligation reaction to do a chemical transformation and it worked (~50 clones per plate).
The T4 ligase is new.
Is it possible for something to happen to my ligation reaction if it sits too long? I did the chemical transformation the next day, but the remainder of the ligated plasmid sat in the fridge for a week before I tried the electroporation.

I havent been doing any ethanol/isopropanol precipitations at all. The only clean up I have done has been on the PCR product, the insert, after PCR.

#4 Knights

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Posted 12 August 2010 - 11:32 AM

I actually had a similar issue recently. We had one type of Quick Ligase kit and then received a kit from another lab. The new kit wasn't working and I was getting 0 colonies when ligating. Unfortunately we didn't have any of the old ligation kit to test if it was a ligation issue or something else.

When we read the kit's manual carefully we found the issue. The new kit's buffer has PEG, which is apparently bad if you're using electroporation.

So while for the old kit we would do ligation --> heat inactivation --> electroporation and it would work like a charm, with this kit we had to do ligation --> ice incubation --> butanol extraction --> electroporation.

The butanol extraction gets rid of the salts and PEG, so electroporation can occur without a hitch.

Lesson learned for us. Hope this helps? Good luck! I know how frustrating it can be.

Also, if when you digest are you checking if the vector has linearized properly? Does your PCR product have enough of a hanging sequence for the restriction enzymes to sit on and do their cutting?

Edited by Knights, 12 August 2010 - 11:33 AM.


#5 Mattise

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Posted 12 August 2010 - 12:14 PM

When we read the kit's manual carefully we found the issue. The new kit's buffer has PEG, which is apparently bad if you're using electroporation.

So while for the old kit we would do ligation --> heat inactivation --> electroporation and it would work like a charm, with this kit we had to do ligation --> ice incubation --> butanol extraction --> electroporation.

The butanol extraction gets rid of the salts and PEG, so electroporation can occur without a hitch.


I was thinking it was something in the ligation reaction, but I am unsure if any of this would really prevent electroporation.
Here is what is in the T4 ligase:
300mM Tris, 100mM MgCl2, 100mM DTT and 10mM ATP

I know salts can ruin an electroporation, but 10-20 ul of a 100mM MgCL2 solution into 200ul of cells? Seems like it wouldn't be enough, and I don't detect any arcing.

I think this is probably my next step though, clean up my ligation reaction.




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