cloning in expression vector - (Oct/02/2007 )

hi all

we have cloned and sequenced an antiviral protein gene that we know is also a ribosome-inactivaing protein that inhibits proteinsynthesis in eukaryotes (we do not know if it does the same in prokaryotes). we now have to express this in bacteria, and are using the pet28a for the purpose. the problem is that we are not getting any colonies in the recombinant plates but the colonies are there in the control plate were the bacteria have been transformed with native undigested pet28a vector. the site being used for cloning is ecorI. the vector was dephosphorylated, and the phosphatase inactivated before ligation.

my problem is that i do not know if the ligation has not worked, or if the gene indeed is expressing, but because of the protein product being cytotoxic, is kiling the bacteria.

could i have your suggestions as to how i can find out which of the two is the reason for a totally blank plate.

thanks

viv

The T7 promoter in pET28a should not be active at all in a cloning strain, so the toxicity of the insert should not be an issue. I'd guess that you are having problems because of your cutting, dephosphoylation, or ligation. The first thing I'd do is to try without the dephosphorylation. Can you digest with two enzymes and avoid religation? Tell us exactly what enzymes are used, how the insert is prepared, how the vector is prepared, how much of each is ligated, how you are doing the ligation, how you are transforming, what strains you are using, what resistances you are selecting with. Everything. Volumes. Manufacturers. Cleanups. We can't guess these things, and you're looking for a problem.

thanks siuchi and phage.

the gene was originally cloned by the t/a cloning strategy in promega pgem t easy vector. this vector has an ecor1 site on both sides of the cloning site so that the fragment can be easily taken out with a single digest. the clone was cut out from the pgem t easy by ecor1 digestion, and purified using a fermentas gel purification kit that works very well. pet28a was chosen for expression cloning since the gene would be in frame, and since it has an ecor1 site that could be used for our purpose. the first occasion, we digested the vector with ecor1, did not dephosphorylate it, but gel purified it as such. both the vector and insert were picked up in 10 µl as per the manufactuer’s protocol. The ligation was set up in fermentas t4 dna ligase, with the buffer provided by the manufacturer. a 1:3 vector:insert ratio was used. the reaction was carried out overnight, and the bacteria was e. coli strain jm109. transformation was carried out with the fermentas transformaid transformation kit. selection was by the kanamycin resistance gene.

the control transformation with native vector worked fine, but no colonies for the recombinant one.

we then did the same after dephosphorylating the vector, killing the phosphatase by treatment at 65 c prior to setting up the ligation. all the rest was as above.

again, the control plates had colonies but the recombinant plates did not.

thanks

viv

Let me understand this correctly,
The vector (pet28a) was linearised with EcoRI. The EcoRI cut insert was added. And ligation conducted. No colonies were recovered from the transformation.

Is this is correctly, I strongly urge you to
1) Check that your ligase is working
2) Check that your ligation mix actually has DNA in it.

Who much DNA did you dephosphorylated and for how long? Over dephosphorylation is a bad thing, as you will damage your DNA ends rendering the vector unligatable. Could you writed down your dephos conditions and the amount of DNA unused.

Conduct your ligation again. (using dephosphorylated vector).
Once ligation is completed, run 1/3 of the ligation mix on an agarose gel (using as narrow a well as possible). If your ligations works, you should see high molecular weight bands (larger then your vector). It also checks to make sure you have not lost your DNA.

By using just one digestion site you are going to need to do the dephosphorylation step to prevent the vector from re-ligating itself but make sure you are using the least amount of CIP/SAP and the least amount of time possible. You need to include a negative control here. After dephosphorylation, try to religate the vector. This will show you if the dephosphorylation was effective. I've always re-purified rather than heat inactivation and I'm not sure if that would have any effects. I'm wondering when you say a 1:3 ratio if you are just using concentration or are you actually calculating the molar ratio? Molar ratio (in my experience) is critical. Just divide the size of the insert by the size of the vector, multiply by the amount of vector DNA used and multiply by three. This is the amount of insert you should use but I always set up a 1:2, 1:3, 1:5. Another important thing is the total amount of DNA being put into the ligation. Many people think more is better but really, it's the opposite. I only use 25ng vector.