Electroporation and Plating of M. xanthus transformants

Dialysis

Prior to electroporation of the plasmid into M. xanthus, excess salts from the plasmid extraction need to be removed, as salt will interfere with the current that passes through the sample. You will float a cellulose membrane (pore size 0.025 µm) in a well containing water, allowing for the exchange of water and salt but not plasmid.

1. Fill a 60mm Petri dish 2/3 full with sterile, diH2O. You'll need one per plasmid. Before you add the membranes, ensure that your plates and work area are somewhere that can sit undisturbed for the duration.

2. Using forceps, pick up a cellulose membrane (pore size 0.025µm) ensuring not to touch it with your fingers. As you pick it up, both blue membrane separators should be removed from either side. You may need two sets of forceps to ensure this without touching. Float the membrane on top of the water using forceps (you'll need a steady your hand as the membrane needs to float on the water without water pooling onto the surface). It helps to pick up the membrane at the very edge with the forceps. Try to keep it as horizontal as possible as you lay it on the surface of the water. Let the membrane float for 5 min to thoroughly wet it.

3. Get at eye level with the dialysis membrane and check multiple angles to ensure that no water has pooled on the surface.

4. Pipette 20µl of each plasmid sample into the center of the separate dialysis membranes, using caution not to touch the membrane too forcefully. The plasmid sample should form a droplet in the center and should not run off either side.

5. Gently cover the plate with the lid and let the plasmid dialyze for 20-30 minutes.

6. Pipette the droplet back off of the membrane, again using caution to avoid flooding the surface of the membrane. Transfer this dialyzed sample to a clean, labeled centrifuge tube (labeled with date, plasmid, and the fact that it is dialyzed). Proceed to electroporation.

Electroporation

While heat shock transformation is commonly used in E. coli, some bacterial species are not as easily transformed and require alternate methods. Electroporation is a technique that passes a current through a sample of cells, temporarily opening pores in the membrane and allowing for a more efficient uptake of plasmid DNA than with heat shock methods.

1. Turn on the electroporator so that it can warm up while you prepare your sample.

2. Add 3.0 ml CTTYE growth media to a 50ml sterile flask for each sample you are electroporating. Set aside.

For each sample:

3. Pellet 1.5 mL of an overnight wild-type M. xanthus culture in a centrifuge tube for 2 minutes at 13,000xg. You don't really need to check the density of this first.

4. Decant supernatant (pipetting off any excess) and add another 1.5 mL of the overnight M. xanthus culture to the tube containing the cell pellet. This will just allow us to spin the equivalent of 3mL of cells without needing to set up the larger centrifuge.

5. Spin again for 2 min to pellet the cells, concentrating the sample. Decant the supernatant and pipette off the excess into the waste.

6. Resuspend the cell pellet in 1mL of sterile MilliQ water, pipetting slowly to avoid spillage over the tube. Spin for 1 minute.

7. Decant the supernatant and repeat that wash step two more times for a total of three washes of the cell pellet in water, resuspending for the final time in 40µl of PCR water. This step removes salt and other components of the growth medium that can interfere with electroporation. Remember that cells will not be very happy in water so try not to leave them sitting in the tubes longer than is necessary in this step.

8. Pipette 10 ul of the plasmid into the tube with the cells and stir gently with the pipette tip to mix.

9. Transfer the 50 µl volume into a 1mm gap electroporation cuvette. The sample should be placed down toward the bottom of the cuvette, between the two metal plates.

10. Electroporate at 650V (time constant should display that it took ~5ms).

11. Add 500µl CTTYE broth to the cuvette and VERY gently pipette up and down to mix. Cells will be fragile at this stage

12. Transfer all 550µl into the recovery flask or tube that you created earlier.

13. Label the tube and place in the shaking (125rpm) incubator at 33C overnight.

Plate M. xanthus transformants (Day 2)

M. xanthus motility mechanisms work well on an agar surface, so plating transformants by spreading will result in a lawn rather than discrete colonies with genetically identical cells. Instead, cells must be embedded into a soft agar substrate and poured over the top of the agar plates with kanamycin selection.

[edited 2/16/23]

Prior to starting this protocol, you should have (for each of your electroporated samples): 1 CTTYE plate and 3 CTTYE/kanamycin plates warmed to 33C 4 15mL conical tubes each containing 3mL of molten CTTSA, incubating in a 55C heat block

1. Remove M. xanthus cells from the shaking incubator just before you are ready to begin. Vortex your culture (with the cap in the fully locked position) and then remove the cap and pipette up and down to mix the culture and break up any large clumps of cells.

2. Remove conical tubes containing CTTSA from the heat block and allow them to cool for ~3 min. The outside of the tube should feel warm but not hot.

3. Working quickly, add 200µl of cells from the recovery flask to one tube of CTTSA, vortex gently to mix, and pour the contents of the tube over the surface of the CTTYE agar plates. Gently tip the agar plate from side to side until the CTTSA covers the entire surface. Cover the plate. This will be your non-selective control plate.

4. Repeat this step with the CTTYE+kanamycin plates, adding: 200 µl of cells 500 µl of cells + 500 µl CTTYE nutrient media Everything else in the recovery flask (minus any large clumps of cells)

5. Allow CTTSA agar to solidify for 15 min before inverting the plates, then incubate at 33C for 5 days for colonies to grow. The timing of this is variable–we are actually generating the mutant in this step, so depending on the efficiency of the transformation and the impact of the mutant on the growth of cells, this can take anywhere from 2-5 days.

Monitor over the next week and:

Patch ~10 colonies onto a CTTYE/kan plate and allow to grow overnight at 33C. Based on the growth rate of your potential mutant, the patches might need 2-3 days.

Number the patches that grew, and select one to transfer to a flask containing 12 mL of CTTYE broth. Shake at 33C overnight.