transformation troubles - where to go next? - transformation/ligation (Oct/20/2006 )
Hi everyone,
After 2 weeks of fighting with this project, I have decided to seek out the wisdom of the Forum.
I am doing what should be a simple "cut and paste" ligation and transformation. I have tested the restriction enzymes (they work), and the restriction sites (A-OK). The T4 is new (Fermentas), the Amp plates are new (with 100ug/ml Amp), the comp cells are new (DH5 alpha, Invitrogen). I have narrowed down the problem to, I think, the transformation. After transforming 5 ul of a 20 ul ligation reaction into 50 ul of DH5, then spreading 100 ul on one plate and the (concentrated) remainder on another plate, as well as a background vector reaction, I get very small, suspicious-looking colonies (lots on ALL the plates!). I tried prepping several of these little guys, and got absolutely nothing! Last night I tried the pUC19 control along with my ligations, and this morning I got the same small colonies on my ligation plates but almost nothing on the pUC plates.
I see that Invitrogen has changed the transformation protocol slightly: it says that you should heat-shock for 20 seconds at 42 degrees: I have ALWAYS done the 45 second thing - the new shorter time made no difference anyways. 
If anyone has ANY ideas, please let me know!
Thanks.
i have a suggestion.
Perhaps you should lower the Ampicillin concentration down to 25ug/ml. I have the feeling that your cells are being killed/inhibited before they have time to express the ampR gene.
There is also a conspicuous absence of a recovery period in your protocol. Sound like the quick transformation protocol... I would have a recovery period of half an hour to help the cells along.
Hi perneseblue,
Thanks for the quick response!
I will definitely try the lower amp concentration (although I will fall back on my standard whine of "the regular way SHOULD work!").
For recovery, do you mean the post-heatshock time on ice? I normally do 2 minutes after the 42 degrees, and before adding the LB.
One thing - do you think it makes a difference whether LB or SOC is used for the 1 hour shaking (37 degree) interval?
Cheers!
My mistake.. so you do recover the cell in rich medium (I use SOC) before plating.
I have never tried to see if there is a difference. However I am told (by the local wisdom) SOC is a better recovery medium then LB because it is a richer medium with lots more amino acids. You could go find out and report back with your findings. 
I agree that the transformation step seems to be problematic.
You need to add some controls: vector only plus ligase, cells only (no added plasmid). The fact that your pUC19 control came up blank is very suspicious. Try another tube of supercoiled plasmid. Is your vector dephosphorylated? BAP and CIAP can result in damaged ends if overused due to contaminating exonucleases. Was your DNA exposed to UV light for more than 30 seconds?
SOC is supposed to result in 2-3x as many colonies as LB.
Amp 100 is required for pUC origin plasmids. Amp 25 is adequate for pBR322, pACYC, pSC101 and other low copy number plasmids. Satellite colonies are a risk if you do not use enough ampicillin.
If your insert is toxic, it may help to grow the cells at 30C rather than 37C.
Hi tfitzwater,
I do use the vector only + ligase control, but have not tried the "cells only" approach. The pUC control was the one that came with the DH5 from Invitrogen, and I used it as per the instructions (yes, I am desperate enough to even follow instructions, instead of going via experience! 
).
The vector is treated with CIAP (even though there are sticky ends, I always like to treat them just in case).
From what you guys are saying, it sounds like maybe the SOC idea is potentially good. I will try it again, and hopefully have good news to report!
Thanks.
I always dephosphorylate even when the vector is being cut with two enzymes because there is always a small percentage of molecules that are only single-cut rather than double-cut. Dephosphorylation prevents those vector molecules from religating.
A caveat on dephosphorylation: the most common reason for failure to obtain colonies is a result of adding too much BAP or CIP to the vector prep. These enzyme preparations are difficult to purify and are frequently contaminated with exonucleases and phosphodiesterases. BAP and CIP are routinely used at elevated temperatures because the contaminants are less active at high temperatures. Excess enzyme will introduce contaminating exonucleases and nibble away restriction overhangs. In addition, BAP is more heat-resistant and is difficult to completely inactivate. Precise determination of the concentration of 5’ ends is required. Calculate the exact number of ends: 2 x (g of DNA)/[size in bp x (660 Da/bp) = moles of ends of a double-stranded DNA. Use exactly enough BAP or CIP for the number of ends. Unit definitions may vary according to the supplier. This usually requires a significant dilution of the stock. Excess CIP is also reported to inhibit complete dephosphorylation. Because the byproducts of the reaction inhibit the reaction itself (dephosphorylation generally only proceeds to 95% completion), I then clean up the DNA by adding 10x stop mix consisting of 200 mM EGTA, pH 8.3, 10x TNE100 (1x TE containing 100 mM NaCl) and 10% SDS and heat inactivating at 56-68°C for 15-45 minutes; followed by phenol:chloroform extraction and ethanol precipitation. Then I repeat the entire CIP protocol all over again to dephosphorylate the residual molecules that failed to dephosphorylate the first time. This has proven to be infinitely safer than using too much enzyme.
SAP is thermally sensitive and must be used at 37°C, but lacks the contamination problems of BAP and CIP. Generally, one unit of SAP is required for the complete removal of terminal phosphates from 100 pmol of 5’ ends of DNA. I have switched to the exclusive use of SAP, but supply these notes on the use of BAP and CIP for those who wish to use those enzymes. SAP is active in restriction buffers, so I add it to the restriction reaction if it is performed at 37C. Heat inactivate all the enzymes at the end and then perform one clean up step instead of two.
That's good to know about the SAP - I have heard of it, but not actually used it before. Also, I did not realize that excess CIP could be inhibitive.
One avenue that I was exploring (ie: grasping at straws), whilst trying to get this thing going, was the restriction sites (for the ligation part). NEB says, in its troubleshooting section, that ligation can be difficult when one of the sites has only a 1 base overhang, and that this requires the use of more T4 ligase. Have any of you come across a similar situation?
Cheers!
I humbly disagree with this assessment pertaining to pUC origin plasmids. I have worked with both pUC and pBlueScript using Amp at 25g/m. Both plasmid are high copy number plasmids. And more antibiotic is not necessarily a good thing. The aim should not be the total annihilation of sateliite colonies, but rather stopping the plate before all the amp is degraded from the culture.
Perhaps you should lower the Ampicillin concentration down to 25ug/ml. I have the feeling that your cells are being killed/inhibited before they have time to express the ampR gene.
There is also a conspicuous absence of a recovery period in your protocol. Sound like the quick transformation protocol... I would have a recovery period of half an hour to help the cells along.
Qiagen protocol from PCR cloning kit suggested 100ug/ml of Ampicillin.
I dont really get it sometimes. Isnt it if the amp concentration is too low, there will be no more selective cells to grow on the media right?