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DIY Electrocompetent E.coli

An image on a halo to depict DIY electrocompetent E coli

If you buy competent E.coli regularly, you’ll know that they are pretty expensive.

So the cost of screwing up a cloning or transformation experiment is pretty high in terms of money, as well as your time and sanity!

But you don’t need this extra worry because despite what their high commercial cost would suggest, making good quality competent E.coli is very easy. One morning’s work (with a bit of work ahead of time) is all it takes to make great DIY electrocompetent E.coli prep.

In this article, I’ll describe a protocol for making electrocompetent E.coli that contains a variety of tricks and tweaks that make it possible to routinely get competencies of 1 \times 10^{10}, with a little practice.

I’ll also describe a couple of quality control checks that you can do to validate each prep you make.

This protocol is for making a fairly large batch of cells but can be scaled down easily without loss of quality.

The tips and tweaks are as follows:

1. Keep everything fresh and chilled at all times
2. Wash the cells extensively in glycerol
3. Start with a high volume of cells so that the final competent cell aliquots are very concentrated.
4 Hand-wash the glassware before autoclaving to ensure that no detergent is present

These are all included in this protocol and the original references are listed at the bottom of this article.

The Protocol

1. Streak the strain you wish to make competent onto an LB plate and incubate overnight at the appropriate temperature.

2. The next afternoon, pick a single colony into 10 mL of LB in a sterile bottle and grow overnight in a shaking incubator at 37°C.

  • At this point chill the following in the freezer:
  • falcon tubes or centrifuge pots (see step 5)
  • 1 L of sterile 10% glycerol
  • 35 sterile cryovials, labeled with the strain name

3. In the morning, inoculate 800 mL of LB in a 2 L baffled flask with 8 mL of the overnight culture and grow at 37°C in a shaking incubator.

4. Grow the culture to an OD of between 0.7 and 1.0 at 37°C. This should take around 2-3 hours.


5. Transfer 400mL of the culture to 8x pre-chilled 50 mL falcon tubes (or a suitably sized sterile centrifuge pot). Chill the tubes, and the remaining 400 mL on ice for 1 hour.

6. Centrifuge for 10 minutes at 4500 rpm and 4°C then very carefully remove the supernatant.

7. Pour the remaining 400mL of culture into the tubes and repeat step 6.

8. Add 5-10 mL of chilled 10% glycerol to each falcon tube and gently re-suspend the cells. Then make up the volume in each tube to 25 mL with 10% glycerol.

9. Centrifuge for 10 minutes at 4500 rpm and 4°C then remove the supernatant.

10. Repeat steps 8 and 9 twice times. On the final repeat, pool all of the cells into 1 Falcon tube, centrifuge as before then resuspend in a final volume of 6 mL in 10% glycerol.

11. Leave the cells on ice for 10 minutes then pipette 180 µl into each cryovial and transfer immediately to the -80°C freezer.

12. Keep the remaining cell suspension for quality control checks.

Quality control checks

A batch of competent cells like this is only good if you actually know how good they are so it is worth performing a couple of simple quality control checks.

1. Phage check

Streak 35 µl of cell suspension onto an LB plate and grow overnight at 37°C. If there is no phage contamination, the cells will grow to form a thick, healthy lawn.

But if phage is present, circular clearings will appear or, if there is a very high amount of phage, there will be no visible growth at all.

2. Competency check

Transform 2x 50 µl of the cell suspension with 1ul of an empty plasmid (preferably pUC18) at 0.1 ng/µl. Plate 5 and 50 µl on separate plates with the appropriate antibiotic selection and grow overnight.

Count the number of colonies on the plates and calculate the number of colonies formed per µg of DNA. (e.g. If you obtain 50 colonies on the 5 cm plate, the efficiency is 1 \times 10^8).

Normally, 1 \times 10^8 to 1 \times 10^{10} cfu/µg DNA for standard 3-5 kb plasmids should be easily achievable with this protocol.

Any questions/comments? Click on the link below to discuss this article in the Bitesize Bio Bistro.

For more tips, tricks, and hacks for getting your experiments done, check out the Bitesize Bio DIY in the Lab Hub.


  • Dower, W. J., Miller, J. F., and Ragsdale, C. W. (1988) High efficiency transformation of E. coli by voltage electroporation. Nucleic Acids Research, 16, 6127-6145.
  • Chuang, S. E., Chen, A. L., and Chao, C. C. (1995) Growth of E. coli at low temperature dramatically increase transformation frequency by electroporation. Nucleic Acids Research, 23(9), 1641.
  • Sheng, Y., Mancino, V., and Birren, B. (1995) Transformation of Escherichia coli with large DNA molecules by electroporation. Nucleic Acids Research, 23(1), 1990-1996.
  • Engberg J., Andersen P. S., Nielsen L. K., Dziegiel M., Johansen L. K., Albrechtsen B., (1996) Phage display libraries of murine and human antibody fragments. Molecular Biotechnology, 6, 287-310

Originally published 25 November 2008. Updated and republished 2 February 2015

Image Credit: Anthony D'Onofrio


  1. Ram on February 21, 2018 at 10:54 am

    Hey Nick, thanks for this protocol.
    But at steps 5 and 7, don’t you think that by splitting the culture into 2* 400 mL for the centrifugation, the cells from the first 400 mL remain in the pellet for too long? Wouldn’t that result in a too high cell death from oxygen deprivation? Why not centrifuge the whole culture in one go?
    Also, one of the biggest problems for me in competent cell preparation is the resuspension step. I need to keep them chilled, I need to be gentle and when they are adamant about sticking to the wall of the centrifugation container, I could spend several minutes trying to get them back into solution. Any tips here?

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