I'm beginner in this field of biotechnology and intend to construct knockout strains of gram negative bacteria by homologous recombination. My question is the following.
After inserting the plasmid into the bacterial cell by electroporation, the truncated target gene (KmR cassette inserted) is supposed to recombine with the homologous region in the chromosome.
Is there a way to verify that the recombination has taken place? Will the complete target gene, harvested in the plasmid, not be expressed?
If you're targeting HR to a specific locus and expect a specific recombination product then you should be able to simply PCR and sequence (and/or do a diagnostic digest on) that locus to verify that it has been altered as expected.
Using sequencing the region of genome where homologous recombination is supposed to be occurred, using PCR designing outward primer for both 5' and 3' from your insert and inward primer from genomic region just outside homologous region.
otherwise you can confirm using Southern blotting designing probe of your insert and supposed to be deleted region.
Gene inactivation in your case can be confirmed in several ways. You should note a loss of function of the targeted gene in Km resistant bacteria that have lost the plasmid. If there is no known phenotype related to the gene, its PCR amplification and sequencing will give you information about whether it is inactivated or not.
I am not too familiar with the constructs for recombination in E.coli. I mostly work in yeast, but the general concepts are the same and chromosomal integration/gene deletion by HR is permanently used in Saccharomyces:
The easiest way is to simply use PCR. Make a specific primer for each site of the targeted chromosomal gene, two primer specific for the insert and two specific for the internal (to be replaced) sequence of the target gene. You should then do five PCRs on chromosomal DNA for your transformants:
1. upstream flanking primer + downstream insert-primer
2. upstream insert-primer + downstream flanking primer
3. upstream flanking primer + downstream flanking primer
4. upstream insert primer + downstream insert primer.
5. upstream target internal primer + downstream target internal primer
#1 and 2 will show you that the insert is flanked by the predicted chromosomal sequence
#3 will tell you whether the size of the targeted region changed (as expected if the inserted part is of a different size than the wild type locus - use a wild type control to see the difference directly on the gel)
#4 will show you whether the insert is present somewhere in the cell at all. e.g. false integration at a different locus would show a band here but not in #1+2
#5 will tell you if the target gene sequence is - for whatever reason - still present somewhere in the genome.
Alternatively, you could also just sequence the chromosomal region to verify that the integration looks as predicted. That would be the best way, but also more expensive.
As for the question on expression: PCR set up #5 will give you a good first impression for this. However, the best way would be a Western blot to show there is no protein expressed in the cells or a Southern blot to verify that the DNA sequence is lost. But for a fast first screening of candidates, the PCR approach is easier and faster.
Hope that helps.
Best regards,
Chris
I am naive in this field but from my previous experience I would like to give some comments which may be helpful.
First, you can check by PCR, the fastest and easiest way and then confirmation by sequencing.
Second, If you know the sequence of previous and expected construct, you can check this by restriction enzyme analysis.
Thank you.
Thank you very much for your answers and advices.
My question was mostly about the following:
1) Checking by PCR, how I may be sure the amplification product comes from the gene harvested on chromosome or on plasmid vector?
2) What happened to the plasmid vector once HR has taken place?
3) If vector remains into the cell, due to the intact gene is still harvested in it, is still possible that the studied phenotype be expressed even after homologous combination have taken place?
Thank you
As you could conclude from my previous answer, you need to get rid of the plasmid. Otherwise, you wouldn't be able to select for insertion event since plasmid is also providing Km resistance. Usually suicide plasmids are used for such situations, i.e. those that are unable to replicate in some conditions (e.g. increased temperature etc.). If you are using an established method, the plasmid should have been curable.
Good luck!
With your description I think you use the pKD46 plasmid as vector for recombination. The classic test is the replacement of a gene by the KanR or CmR resistance cassette. To select only you recombinants, you just have to plate you electroporated cell on kan selective media. And if i'm right at 37°C the plasmid is non replicative and you loose it.
To verified you just have to isolate your clone by streak on selective media and sequence the putative replaced gene using primer at the outside of the replacement.
If you discribe more your protocol maybe I can be more precise on how to do this.
Thank you Damir and Geoffrey, very good information.
Actually, the plasmid used as vector is pUC119 (LacZ+AmpR cassette) and pHSG299 (lacZ+KmR cassette).
Transformed bacteria are not E. coli, but acetic acid bacteria from the genera Acetobacter or Gluconacetobacter, and they grow at 30°C.
Are those plasmids suicide vectors or non-replicative?
pUC119 is a high copy number plasmid, so it is very difficult to remove it from a bacteria. Your second plasmid have the same origin of replication, and they must enter in competition each other. I read this second plasmid is an expression plasmid and I not recommend it to do gene replacement. This two plasmids alone are (usually) very stable.
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