For a plasmid as large as this I would just do a tried and true phenol:chloroform extraction and ethanol precipitation. Use a coprecipitant such as glycoblue and wash the pellet carefully to remove salt. I have used this prior to electroporation with good results.

Alternatively, NEB now sells a ligation mix that can be used directly for electroporation without cleanup.

For electroporation, I just do Ethanol Precipitation without additional cleanup. Just add 1/10 volume of 3M Na-Acetatete (pH 5.2) and 3 vol pure Ethanol, mix and incubate 5 min at -80°. Spin 10 min max Speed (16000rcf), wash once with 1ml 70% Ethanol to remove Salt, take off supernatant completley and resuspend DNA in 5-10µl of H2O, rinsing the wall of the tube.

Alternatively, up to 1µl of unpurified ligation usually can be electroporated without arcing (helps to lower the voltage a bit, I use 1350V instead of 1800V on a GenePulser XCell).

Good Luck!

Best regards,

Thomas

I agree with Arjan. Purifying the plasmid post-ligation seems unnecessary. I would simply try cells with high competency. Also, most ligation reactions work best with total DNA in the range of 20-100ng. Several hundred nanograms may be detrimental- why do you need so much?

@ Andrew Tai: Thanks for the suggestion, I am hesitant to use phenol:chloroform extraction as in the past that I have done this, I've gotten nicks introduced through the vortexing and harshness. It's probably not even necessary for purifying this reaction. I will definitely try out glycoblue+ethanol precipitation.

@Thomas Zillinger: Good to know that you can do ethanol precip with Na-Act is compatible with electroporation. I assume you are using 1350v on 1 mm cuvettes? I've been doing 1500-1800 under the assumption that you want the highest voltage with acceptable cell death. In your experience is a better lambda closer to 5 more beneficial than a higher voltage?

@Arjan Groot and Daniel Cohen: One of the pieces prior to this final construct was a 15kb plasmid which electroporated quite well, but gave me no colonies chemically transformed. I figure that 15 kb has traditionally been a barrier for chem compt cells. I gather you guys have had success chem competent transformations with large plasmids?

In addition, in a, perhaps failed attempt to simplify my question, it's not so much a ligation as an in-vitro assembly reaction. In order to reach .02 pmol/fragment in the reaction and because the fragments are in the kb range, it takes quite a bit of dna to reach .02 pmol. It would be interesting to see if the ligation rule of less is usually better works for this as well, since it operates on similar mechanisms.

I have been hoping that the reason why I'm getting no colonies is because yield of circular dna is too low. Papers have examined the role of DNA concentration on electroporation efficiencies, and it seems to be a linear relationship over 2 log ranges. Thus, I am hoping that concentrating my 20 uL rxn into 1 uL will yield an increase of colonies...from 0 to >0. *hopefully.

But it's a good idea to try lower dilutions.

Thanks guys for all the suggestions and for being my sounding board!

Brian

Brian- are you assembling PCR generated fragments?

@ Karen Posey: That...is a very interesting technique! It's like a DIY osmotic purification. What is the purpose of 10 mM Tris and not di-water? Also, do you happen to remember a rough material/pore size for the membrane? I have 45 and 22 um filters in PVDF, PES, but I assume since people blot into PVDF it might not be the..best idea.

@Daniel Cohen: I am assembling a medley of PCR generated fragments as well as digested pieces

Brian- I think you could get this to work if you increase the efficiency of fragment hybridization. One approach is to use Gibson assembly, another is to use restriction enzymes such as SapI that allow you to perform a single digestion for all your pieces but specify unique 3bp overhangs that yields directional assembly. I would expect that either approach would greatly simply your assembly reaction and yield a higher number of clones than your current set-up.

@Daniel Cohen: Oops, sorry for the long delay. I thought I had responded and realized I hadn't hit enter!

I actually am using Gibson assembly at the moment as my construct needs seamless construction.

Unfortunately, it appears there are a large number of variables which affect gibson efficiencies. I have doubled overlap lengths to 40 bp.

Have you had any experience with Invitrogen's GeneArt enzyme mix? Their support claimed it was a different mechanism to gibson assembly, and they have much more troubleshooting/documentation on the process.

Thanks

Brian

Hi Brian. I haven't tried the GeneArt enzyme mix. We've used Clontech's In-Fusion kit with solid results. I've also done multi-site recombination with LR Clonase II, but you're stuck with the att sites which may not be suitable for your purposes. I think high DNA amounts may the culprit in your system. This might be somewhat unfavorable for the Gibson assembly, but is highly unfavorable for successful transformation. Have you tried diluting your sample 1:20 and using an ultra high competency cell type for the transformation step?

I've since had a brief talk with an NEB specialist about it, and he has assembled a 18kb construct with 20 bp overhangs, but by loading an absurd 300 ng of backbone, and 1:1 molar ratios of the remaining pieces. He did dilute it 1:3 prior to electroporation. It does seem counterintuitive that higher amounts are necessary. In my mind, it would just sequester the enzymes and not distribute them properly.

A very interesting tidbit came out of that talk though, that he has not been able to get gibson assembled reactions to transform with invitrogen competent cells, but has gotten them to work with straight up DH5alphas, Sures, and of course, NEB cells. I just got in some of their 10beta cells, so I will actually just try electroporating the stuff I have leftover.

I have only diluted 1:3, and gotten zero colonies. 0 background is very nice since I'm doing dual selection of Cam/Kan in the backbone/insert.

What do you consider to be ultra high competency? I've thus far tried electrocompetent Top10s, Stbl4s, and soon to be 10beta.

Brian- I've had good success with Agilent (formerly Stratagene's) high competency cells: http://www.genomics.agilent.com/CollectionSubpage.aspx?PageType=Product&SubPageType=ProductDetail&PageID=643

XL10 Gold Ultracompetent have never failed me, yet!

Maybe you will fare better with heat shock than electrocompetency, since I wouldn't expect it to be as sensitive to salt levels.

It is curious that you mention that you have Cam resistance in addition to Kan. Is it possible that your Cam gene isn't being expressed constitutively? Any chance there are any toxic genes in your plasmid? Either phenomenon would lead to zero transformants, irrespective of recombination and transformation efficiencies. Depending on your plasmid design, you might do better to initially select for KanR, and then replica plate for CamR.

Thanks for the XL10 recommendation, I have heard of them amongst the list of high efficiency cells, but never had any particular reason to single them out.

There are multiple reasons I can see heat shock being better than electrocompetency, such as salt levels, introducing PEG, exonuclease, ligase into a cell via electro, however I had originally thought that it would be counter balanced by the fact that its a fairly large construct.

I'm transforming the 10betas by heat shock right now, so...we'll see how that goes soon. *crosses fingers

Ahh, yes the Cam+Kan issue. I have definitely lost sleep over that problem. In my final rationale, the Cam is part of the original minimal backbone. I have since manipulated that to add more sequences to it, and it continues to function, so I am fairly confident that is working.

The Kan is driven by the EM7 promoter, and works in the plasmid it comes from as well. Of course, what would be instructional would be to make a backbone(CamR)-EM7(KanR) construct and grow it on my plates to check...but with all the projects I have running, that particular control has kind of slipped under the radar/motivation. I will probably kick myself later when I do generate that control though if it turns out there is a problem somewhere there, and not with the synthesis.

I am intrigued by sequential selection though. I am setup for replica plating as it is, so perhaps that is worth a try with the transformants I have going now.

Again, thank you for all your advice thus far, it has been invaluable.

Brian