Hi,
I'm trying to disrupt some genes of the genome of E. coli. For achieving this, I'm following the protocol of the P1 phage in which this phage transfers genetic material from one strain to another. The recipient strain is from the keio collection, so I don't have any problem with the selection of the colonies which were succesfully trasduced.
But after the confirmation of the disruption with the antibiotic screening and PCR, I tried to remove the Kn resistance using flippase. The flippase is a recombinase that recognize FRT (Flippase recognition target) sites and removes all the flanked area.
This plasmid must be electroporated and then grow it at 30°C, because it has a temperature sensitive ori.
I have done all of this but I have not obtained any transformant. Do you have any clue of why I don't get colonies? Can you give me some tips?
Hi, Mercedez!
Usually electroporation is a much more effective transformation method than the normal heat shock transformation, therefore you might be experiencing problems with your electrocompetent E. coli or the electroporation protocol. Have you tested your protocols before? You should also make sure that everything with your plasmid is OK before you start optimizing your electroporation. The easiest way to do this is to make a restriction analysis and/or send your plasmid for sequencing. However, if you think the problem lies in the electroporation, I can recommend you the following protocol, it worked very well for me:
Preparation of electrocompetent E. coli cells:
Innoculate 3 ml SOB (or SOC) liquid medium with a fresh single colony of your strain of interest and incubate overnight at 37⁰C with shaking. Use 200 μl of the overnight culture to inoculate 50 ml SOB (or SOC) medium. Grow cultures at 37⁰C with shaking to an OD600 = 0.6-0.7, afterwards incubate cultures on ice for 1 hour. Transfer cultures to 50 ml sterile, pre-chilled centrifuge tubes and centrifuge them at 1000 x g for 15 minutes at 4⁰C. Resuspended the cell pellets were in 50 ml cold (4⁰C) sterile water and centrifuged at 1000 x g for 15 minutes at 4⁰C. Resuspended the cell pellets in 25 ml cold (4⁰C) sterile water and centrifuged at 1000 x g for 15 minutes at 4⁰C. Resuspended the cell pellets in 2 ml cold sterile 10 % glycerol and centrifuge samples were centrifuged at 3000 x g for 15 minutes at 4⁰C. Resuspended the cell pellets in 200 μl cold sterile 10% glycerol. You can either use the cells immediately for electroporation or store them at -80⁰C. For storage the electrocompetent cells should be incubated 1 additional hour on ice and stored as aliquots of 40 μl at -80⁰C.
For electroporation:
If using stored cells, let them thaw slowly on ice. Add 1μl (1- 100 ng) DNA to 40 μl electrocompetent cells and incubate 10 minutes on ice. Transfer the reaction mixes to 1 mm pre-cooled electro cuvettes. Use an electro shock of 1.8 V for 3 seconds. Immediately add 1 ml SOC medium to each cuvette, transfer the whole volume of the samples to culture tubes and incubate at 37⁰C (or appropriate temperature) with shaking for 1 hour. You can plate various amounts of culture (I usually dilute my cultures 1:10 and plate 100 μl of the dilutions) on agar-containing selective media and incubate overnight at the desired temperature.
Best of luck!
I plated my cells in LB/Cb (50ug/mL).
Thanks for the recommendation Dilyana, I will try your method.
I initially had problems with this vector too. My solution was to plate on LA-Cb (25 ug/ml) and perform phenotypic expression for 2 hours at room temperature and finally incubate plates at room temperature. This might give some background on the negative control plate (more or less depending of how long you have stored your LA-Cb plates). This can be solved by patching clones on plates with a higher concentration of Cb. This vector is VERY sensitive so anytime i go beyond 25 degrees i start to see a reduction in transformation efficiency.
I should add that i always transform using a modified version of the Hanahan protocol, which is a chemical transformation.
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