Making mutants in Dickeya and Pectobacterium species

Objective: To disrupt a gene in a soft rot bacterial genome.

Insertion mutation: an antibiotic resistance gene cassette is inserted into a convinient restriction site in the gene. One major pitfalls with this type of mutant is that the remaining N-terminal portion of the protein is sometimes functional and can even have dominant negative effects.

Deletion-insertion mutation: the entire coding region of the gene is replaced with an antibiotic resistance gene cassette.

When possible, it is best to make two independent mutants concurrently since on occasion random secondary mutations with interferring phenotypes can occur.

Step 1. Clone the gene or flanking regions into a plasmid

Insertion mutation: PCR-amplify the gene fragment and clone it into pGEM-T Easy or another convinient plasmid vector. Insert an antibiotic resistance gene cassette into an internal restriction site using standard cloning methods.

Deletion-insertion mutation: PCR-amplify the regions bordering the gene of interest. Design your cloning strategy to allow for insertion of an antibiotic resistance gene cassette. For example, cross-over PCR may be used to amplify the gene borders and add a convinient restriction site. Alternatively, the two borders may be cloned with convinient restriction sites and assembled in pGEM-T Easy or another plasmid vector.

When it works, cross-over PCR is fastest.

When all else fails:

PCR-amplify the left border with primers that have XbaI (forward primer) and BamHI (reverse primer) sites added. Amplify the right border with a forward primer that has an XbaI site and SmaI site. Amplify the resistance gene cassette with primers that have BamHI and SmaI-BamHI sites added.

Clone all three fragments into pGEM-T Easy. Then insert the the resistance gene fragment into the BamHI sites of the left border clone. Finally, insert the left border-resistance gene construct into the XbaI-HindIII site of the right border clone.

Primer Site Amplified Fragment Primer Site
XbaI Left Border BamHI
BamHI Kanamycin Resistance Gene HindIII – BamHI
XbaI – HindIII Right Border none

Final Construct:

XbaI Left Border BamHI Kan Resistance HindIII Right Border


Step 2. Electroporate the plasmid into Pectobacterium or Dickeya

  1. Streak the bacterial culture on to LB agar and grow it overnight at 36C.
  2. Transfer a few loopfuls of bacteria to liquid LB medium and grow at 36C for four to five hours. (Alternatively, start a overnight culture, then make a 1:100 dilution and grow the culture for four to five hours at 36C)
  3. Harvest cells from 1.5 ml of culture by centrifugation for 1 min in an epi-tube at high speed.
  4. Wash the cells three to five times with cold filter-sterilized 10% glycerol. After the final wash, suspend the cells in 400 ul of 10% glycerol and add plasmid DNA. Transfer the suspension to a chilled cuvette.
  5. Electroporate at 2.5 kV, 25 uf, 400 ohms, in a 2 mM cuvette.
  6. Immediately add 800 ul of SOC medium. Transfer the mix to a sterile plastic 4 ml tube. Let the cells incubate on the bench top without shaking for at least one hour. Plate cells on LB plus appropriate antibiotics.

We have found that additional washes or adjusting the DNA concentration (less is usually better) affects transformation efficiency, while adjusting the electroporation values (kV, ohms, etc.) does not increase efficiency.

The cuvettes may be rinsed several times with water, then 70% ethanol, autoclaved, and reused several times. The covers can not be autoclaved, but are not required for successful electroporation.

Step 3. Allelic-Exchange.

The goal of this step is to screen for strains where a double cross-over has resulted in an exchange of the mutated gene from the plasmid onto the chromosome.

  1. Choose two independent transformants and start liquid cultures in LB plus antibiotic (for example, if you have inserted a kan resistance gene cassette, use kanamycin. For the rest of this protocol, kanamycin is used as an example). Grow the cultures overnight in LB at 36C.
  2. Wash cells with 100 mM phosphate buffer (pH 7.0) two times to remove the LB medium.
  3. Inclubate the cells overnight at 36C in 100 mM phosphate buffer plus kanamycin.
  4. Sub-culture the cells into LB strains daily by transferring 10 ul of the old culture to 3 ml of fresh LB plus kanamycin medium.
  5. After two or three subcultures (two or three days), dilution plate the cultures onto LB plus kanamycin. Pick colonies to a grid on LB plus kanamycin and LB plus ampicillin. Colonies that only grow on the kan plates have lost pGEM-T Easy and have the kan gene presumably inserted into the genome. Generally, only 100 to 200 colonies need to be screened. If no mutants are found, go back to step 2 and incubate again in phosphate buffer.
  6. Confirm mutations by PCR and/or DNA hybridizations.



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