The Best Way to Desalt DNA for Electroporation |
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Nick Oswald |
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After ligation, the method you use for desalting your sample prior to electroporation is critical, especially if your ligation is inefficient, according to a study by Schlaak et al [1].
Under standard electroporation conditions, the electric field of 12-18 kV/cm generated in a 0.1mm-gap electroporation cuvette means that the conductivity of the sample must be kept very low to prevent arcing. This means that any more than a very small amount of ligation mixture added to the competent cells will cause the sample to arc and the electroporation to fail.
So Schlaak et al looked at various methods of de-salting the DNA and compared how well the purified samples performed in electroporation. And their results were quite illuminating.
The experimental set-up was pretty simple. They took 1ng (high conc) or 3pg (low conc) intact pUC19 in 100ul of ligation reaction mix and desalted the samples using either ethanol precipitation, gel filtration, drop dialysis or a commercial microcolumn (the type used for gel extraction/PCR cleanup), resuspending in a final volume of 50ul.
Then the resulting sample was used to transform 50ul of electrocompetent E.coli and the number of resulting colonies counted. Each experiment was performed in triplicate.
The Results…
As the table shows, for the high concentration samples the microcolumns were the clear winner, although dialysis and gel filtration gave just a 2-fold lower efficiency.
But the difference was more marked with the low concentration samples. Again, the microcolumns were the best, but this time they were at least 100x more efficient than any of the other methods.
For some reason, ethanol precipitation, a widely used method of desalting DNA, gave miserable results for both samples. It’s not clear whether this effect only occurs in the hands of these authors, or whether using a carrier would have helped (they didn’t). But it is certainly worth noting.
The take home message
For desalting 1ng or 3pg of intact plasmid, commercial microcolumns gave superior transformation efficiencies compared with the other methods.
The effect was far more obvious with 3pg of plasmid, which is worth noting, because many of the ligations you do will have DNA concentrations in this range.
So maybe using microcolumns for desalting your ligation mixes could improve your results…
If you give it a try, let us know how it goes.
Reference
1. Schlaak et al Biotechnology Letters (2005) 27:1003-1005
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David Schoppik
Wouldn’t the purification kits add guanidine salts that weren’t present in the ligation mixture (i.e. Qiagen buffer PBI)? I’m not familiar with the kit that the authors cite, but even with thorough washing, in my hands, it is difficult to entirely get rid of those salts (measured by 260/230 spec ratio), especially for small amounts of DNA.
Nick
Hi David,
I thought that too. I think the kit they use in the publication is fairly standard so you would certainly expect some guanidine salts to carry through.
Wendy
I am not convinced by this paper!
I have a problem with the protocol for ethanol precipitation.
We have had good luck using 1 ul glycogen (20mg/ml) as a carrier and Ammonium acetate as the salt (Bring volume of ligation to 99ul and add 50 ul of 7.5M NH4 acetate, 1 ul glycogen, 350ul EtOH) Spin, Wash with 70%, dry. Residual Sodium acetate as they use would cause arcing (ammonium ion would evaporate, sodium would not).
Columns would be very expensive to use routinely especially if one does a careful experiment with lots of controls.
Kurt
There’s also the simple method to desalt the samples in agarose slots, microcolumns are not good for bigger plasmids:
Atrazhev AM, Elliott JF. Simplified desalting of ligation reactions
immediately prior to electroporation into E. coli. Biotechniques. 1996
Dec;21(6):1024. PubMed PMID: 8969827
Nick
Wendy: Using and ammonium salt so that the ion evaporates is a very interesting idea.
Nick
Kurt: Have you tried that method? I always thought that these agarose slot approaches were quite fiddly. How did you find it?
max
Hm, I don’t understand why anyone would still electroporate … didn’t you write some months ago that there is no need today to electroporate, with more competent bacteria?
Nick
… it depends on the situation…!
Shaun
Has anyone worked out if you can reuse the microcon filters? I’m about to give it a go. I’m a bit worried about the stability of the regenerated cellulose filters but they can handle 1 M HCl (enough to break up the DNA) then I’ll run MilliQ a couple of times. I’ll submit the retentate for genotyping and I’ll run a pcr on it too.
Nick
Hi Shaun,
Cellulose is pretty tough. I would imagine that they would survive your washing. Let us know how your experiment goes — it will be valuable information!
vflorelo
Here is a rather cheap and easy and quick way to desalt your plasmid preparations before electroporation.
You don’t actually need to use those fancy filter units, or agarose plugs or anything.
The first thing you need, is a 0.02 micrometer pore membrane, millipore membranes work pretty well.
Second, you need a clean non-sterile petri dish, glass or plastic, no difference
Third, Deionized water
Fourth, well… your plasmid.
Fill the petri dish with deionized water, then place the membrane above the water so that it can float freely.
Then, put a 10 microliter drop of your plasmid preparation on the membrane, and let it stand for 10 to 15 minutes, the membrane pore is small enough not to let DNA pass through (even small plasmids don’t get through it).
Since the drop has enough surface tension for not getting absorbed to the membrane, you can easily recover your 10 microliter drop, free of salts that can screw your transformation.
Another tip, is that you can put more than 10 samples on a 25 mm diameter membrane, only make sure to mark where you put each sample.
After this rather simple procedure your plasmid is really ready for electroporation.
I’ve tried this method, and made very large libraries without crashing the electroporation cuvettes, yielding about 50,000 clones per cuvette (about 3 ml of transformed cells)
Lestress
That’s Hot, thats all!