Electroporation of electrocompetent cells of E. coli or Pseudomonas
Equipment and Consumables:
Sterile workspace (very important for this experiment)
Ideally perform within a laminar flow hood. Under a flame is fine if you have excellent technique.
Incubator (or) Hot Water Bath, ideally shaking at 37°C
Electroporation cuvette
Electroporation machine aka. Electroporator
5-15 ml Sterile LB Liquid medium
Electrocompetent cells of your desired strain (kept at -80°C)
Sterile LB Agar plate containing correct concentration of antibiotic corresponding to plasmid being inserted
Antibiotic Stock Solution corresponding to the plasmid being inserted
Positive control plasmid (shares antibiotic resistance with experimental plasmid)
If you’re just inserting a known plasmid in order to make more stock, a positive control plasmid is unnecessary.
Experimental plasmid, such as a Ligation reaction or Miniprep sample
Ice box with ice
Protocol:
Day 1:
Retrieve the appropriate number of cell aliquots from the -80°C freezer and allow to thaw. You can do this by putting them directly on ice and waiting a long time (~20 min?) or you can put them in room-temperature water in a floatie, then put on ice as soon as they are fully thawed (~5 min?). In the latter case, don’t leave them too long at room temperature, keep an eye on them ! Occasional mixing by flicking helps to speed the thawing process.
<<<note ! don’t try to electroporate chemically-competent cells, these will spark! >>>
Divide up the cells into multiple 50 µl aliquots in sterile 1.5 ml Eppi tubes, on ice. Also put on ice the same number of sterile electroporation cuvettes (2 mm gap size). These need to chill for at last 10 min before use. Label the cuvettes on the cap before use.
Add your plasmid DNA or ligation mixture to the cells. For purified plasmid, 1 µl is heaps ! (and may even be too much). For ligation mixture, it would be typical to use 2-3 µl. In both cases more is not better, because the more DNA you add, the more salt you are also adding, and it doesn’t take very much salt to make the electroporation procedure fail (short circuit = sparks!). Make sure you change pipette tips for each DNA sample.
For all electroporation experiments, you should include a negative control (no DNA added) – this will let you know firstly whether your competent cells are OK (not too salty), and secondly whether your aseptic technique is good. If your negative control makes a spark, your cells are no good (need more wash steps!) or the electroporation machine is not set up correctly. If the electroporation procedure works (no spark), but you get colonies on the plates from the negative control, this means you have contamination, either in your competent cells, or getting in somewhere in the electroporation procedure.
In the case of electroporation experiments with ligation mixtures, make sure you also include a positive control, which is 1 µl of a purified plasmid of known good quality with the correct antibiotic resistance. This should give you thousands of colonies or a confluent lawn of growth on the appropriate antibiotic agar plate – if it doesn’t, this could mean your cells are not competent, or the plates are bad, or the electroporator is not set up properly, or your ‘good quality’ plasmid stock is no longer good quality.
Once all your cell aliquots have the appropriate DNA samples added, transfer each mixture individually to the corresponding electroporation cuvette (again make sure everything is labelled before starting!). Give each cuvette a few taps on the bench to make sure the cell mix is at the bottom of the cuvette.
Take your esky, a p1000 pipette, box of blue tips, some sterile LB broth, and some paper towel over to the electroporation machine (You need ~1 ml LB per sample). Turn on the electroporator, and choose “Exponential” protocol. Adjust the settings on the electroporator to 2500 V, 25 µF, and 200 W. These settings will work with a wide variety of Gram-negative bacteria, but note that for Gram-positives you need to increase the ohms to 800 W (see next protocol). Also note that if you use cuvettes with a different gap size you need to adjust the electroporator settings.
Take your first cuvette off ice, and wipe down the outside and underneath briefly with paper towel. Place this firmly into the cuvette holder on the machine, then close the lid.
Press the “Pulse” button. After a few seconds, the machine will beep to let you know the pulse has been delivered. Note down the time constant displayed on the screen (ms= milliseconds). This tells you how long the pulse lasted, and a higher number here is better. Retrieve your cuvette and immediately add 1 ml of sterile LB to it. Return to ice.
Repeat steps 8 and 9 for all your samples. When finished, turn off electroporation machine, and return to your bench.
As the sample becomes more salty with increasing amounts of DNA, the time constant will decrease until eventually you will get a spark (short circuit). If any of your samples make a spark, this means either your competent cells are not prepared properly (the negative control will spark too), or that you have added too much DNA and/or the DNA is of low quality. It is possible to sometimes get transformants from a cuvette which as sparked, but don’t count on it! Its better to repeat the experiment using less DNA (or a dilution of the DNA in sterile MQ water) and/or a different batch of electrocompetent cells.
Aseptically pour each transformed cell mixture into a sterile 1.5 ml Eppi tube, as follows. First, label all the Eppi tubes, then open them all, positioned in a rack.(they don’t need to be in ice). Take the cap off the first cuvette and briefly pass the top of the cuvette through the Bunsen flame a few times (if you flame it too much it will melt!), then pour into the first Eppi tube. Tap the cuvette on the tube to ensure you get all the sample out. Repeat for other tubes.
Ensure all the Eppi tubes are tightly closed and labelled, and then incubate with shaking for 1 hour at 37°C (E.coli) or 1.5 hours at 30°C (Pseudomonas). Put the tubes lying on their side in a beaker or other container in the shaker, or you can attach them to the shaker with elastic bands or masking tape. Lying horizontal is important for good shaking action in these small tubes.
The role of this ‘recovery’ step is to allow the cells to create the proteins required for antibiotic resistance.
Retrieve tubes from shaker. Prepare 2x the number of appropriate agar plates as you have samples (double check you are using the correct antibiotic(s)!!!), and label all these plates before going any further. In addition to the usual information (antibiotic, plasmid, strain, date), label half of the plates “100 µl” and the other half “pellet”
Spread plate 100 µl of each sample onto the appropriate antibiotic agar. Centrifuge the remaining culture (2 min at 10,000 g in Eppi centrifuge), pour off most of supernatant, and resuspend the pellets by vortexing in the drop of liquid that remains. Plate these resuspended cells on a second set of plates of the same antibiotic type. The reason for doing two plates for each sample is to give us the best chance of getting countable and well-isolated colonies on at least one of these plates.
Also be sure to correctly plate your positive (known plasmid added to cells, plated on antibiotic plate) and negative control (no plasmid added, plated onto antibiotic plate).
Incubate overnight at 37°C (E.coli) or 1.5 hours at 30°C (Pseudomonas). If no colonies appear on any plates, you may have to wait several days.
Don’t throw out the used electroporation cuvettes. These can be recycled. (repeated rinses with water then 80% ethanol, see elsewhere for this protocol). Don’t let the cuvettes dry out with cell mix still in them, you should give these a couple of rinses with WATER as soon as you finish the experimental part of the work. (discard rinsate into culture waste).
Day 2:
Once you have clearly grown colonies, it is time to interpret the results. When examining your plates, first check your controls. The positive control should have thousands of colonies, perhaps even a confluent lawn of growth, especially on the ‘pellet’ plate. the negative control should have no colonies at all. If you don’t see these results with the controls, anything you see on your experimental plates is questionable.
Common problems and their interpretations:
Problem: No growth on your experimental plate or negative control, but plenty of growth on your positive control.
Your Electrocompetent Cell Protocol and Electroporation protocols are fine, something is wrong with the experimental plasmid.
Proceed to Ligation Troubleshooting
Problem: Lots of growth of the negative control (thousands of colonies or lawn). Interpretation:
Forgot to add antibiotic to the plates
Antibiotic concentration is wrong (too low)
Host bacteria are already resistant to the antibiotic (e.g. TOP10 has chromosomal streptomycin resistance)
Plates incubated too long (especially with LB-ampicillin)
Severe contamination with an antibiotic-resistant bacterium (not E.coli) (unlikely!)
Mix up of labelling somewhere – is this actually the positive control? or one of the experimental tests?
Problem: Some growth on the negative control (a few colonies). Interpretation:
Contamination during the procedure, e.g. from one of the other samples or the pipette etc. This may not be a ‘deal-breaker’ so long as there are lots more colonies on your experimental test plates
Antibiotic plates might be old, consider pouring fresh ones.
Mix up of labelling
No growth or very little growth on the positive control plate. Interpretation:
The cells are not competent
Used the wrong antibiotic in the agar (check the sequence of your plasmid to confirm correct resistance)
Used the wrong concentration of antibiotic (too much)
Agar plates are ‘bad’ for some other reason (e.g. added mercuric chloride instead of sodium chloride!)
Plasmid stock has gone bad (run a gel to check)
Mix up of labelling
Pipetting error (look at the pipette tip to ensure that you really have 1 µl of plasmid in there!)
Acknowledgements:
Coleman Protocols 2017 + 2019 http://coleman-lab.org/