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DIY Electrocompetent E. coli

Posted in: Protein Expression and Analysis
Women holding a broken electricity cable looking like she's been electrocuted to represent the competency of electrocompetent E. coli

If you buy electrocompetent E. coli strains regularly, you’ll know that they are pretty expensive—in part because they need to be shipped on dry ice.

So the cost of messing up a cloning or electroporation 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 electrocompetent 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, you’ll discover a protocol for making DIY electrocompetent E. coli strains that you can use for electroporation. I’ve also included a variety of tricks and tweaks that make it possible to routinely get competencies of 1 \times 10^{10}, with a little practice.

You also learn 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.

Tips and Tweaks

Before you get started making your bacteria batch, there are a few things to remember:

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. (1-4)

The Protocol

1. Streak the strain you wish to make competent onto a Luria broth (LB) agar plate and incubate overnight at the appropriate temperature.

2. The next afternoon, pick a single colony into 10 mL of LB medium 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 growth media 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 OD600 of between 0.7 and 1.0 at 37°C. This should take around 2-3 hours.


5. Transfer 400 mL 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 400 mL 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 two more 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 \mul 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.

Phage Check

Streak 35 \mul 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.

Transformation Efficiency Check

Before we explain how to check transformation efficiency, let discuss what transformation efficiency is and why it’s important.

What is Transformation Efficiency?

Transformation efficiency is a way of measuring how well the competent cells take up the DNA during bacterial transformation. It is expressed as the number of colony-forming units (CFUs) that are produced following transformation with 1 \mug of plasmid DNA.

The higher the transformation efficiency the more clones you will get, so it’s important that the competent cells you make have a decent efficiency. Too low and you risk getting only a few or potentially no colonies.

How Do You Check The Transformation Efficiency of your DIY Electrocompetent E. Coli?

Transform 2 x 50 \mul of the cell suspension with 1 \mul of an empty plasmid (preferably pUC18) at 0.1 ng/\mul.

Plate 5 \mul and 50 \mul on separate plates with the appropriate antibiotic selection and grow overnight.

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

Normally, an efficiency of 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.

Tips to Maximize You Transformation Efficiency

  • Avoid using cells that have gone through repeated freeze-thaw cycles. Ideally, freeze in aliquot sizes that you will use in one sitting.
  • Adding 10 mM \beta-mercaptoethanol to growth media and 0.03 mM in transformation media prior to electroporation has been shown to increase the transformation efficiency of competent cells. (5)
  • Use a recovery medium—the process of electroporation is stressful to cells. You can aid their recovery from the stress of perforation by using a high-nutrient medium such as SOC medium following transformation.
  • Purify DNA before transformation, as salts and other contaminants such as ligase can inhibit the transformation process. You can use spin columns such as those in miniprep kits to purify your DNA.
  • E. coli like warm conditions—if you store your plates in the fridge, prewarm them to 37°C before plating out.

Have you used DIY electrocompetent E. coli for your transformations? Let us know your tips and tricks in the comments below.

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

References and Further Reading

  1. 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.
  2. 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.
  3. 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.
  4. 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.
  5. Janjua, S., Younis, S., & Deeba, F. & Naqvi, SMS. (2013). High efficiency DNA transformation protocol for Escherichia Coli using combination of physico-chemical methods. International Journal of Agriculture and Biology. 16:1560-8530.

Originally published 25 November 2008. Reviewed and updated September 2021.

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  1. Maciej on January 24, 2018 at 10:26 am


    thanks for the article. Today I am preparing electocompetent E. coli cells with this protocol. Alas, wilde-type strain. I hope it works! Anyway, common practice in one of the labs I worked in was to put vials directly into liquid nitrogen before moving them into – 80 C. What do you think about it?

  2. Salma on July 6, 2013 at 9:18 am

    Thank you for the interesting protocol, Would like to see the effect of these modifications to the regular protocol.

    I have a question on this part please “Count the number of colonies on the plates and calculate the number of colonies formed per ug of DNA. (e.g. If you obtain 50 colonies on the 5ul plate, the efficiency is 1×10^8).”

    Could you show me how you calculated it?
    (I never bothered to calculate efficiency, either it worked or it didn’t)

  3. Vinu on December 26, 2012 at 2:09 pm

    Hi Nick
    I know your protocol is best for E. coli DH10B and other competent host
    But I am looking for a 10^8 or 10^9 efficiency in MG1655.. Could you help me with this one.. I usually get 10^6 or 10^7 but I really want 10^8 or 10^9

    • Sinke on July 5, 2013 at 10:31 am

      I’m trying to do the same thing. It seems most protocols for preparing competent cells (chemical or electro) are optimized for DH5a and similar strains, so they don’t work so well on wildtypes

  4. owenchinaxiong on March 11, 2012 at 11:25 am

    Is a 15ml falcon tube required? Is it ok to use an EP tube instead? Thanks!

    • jkimmey on March 15, 2012 at 2:34 am

      You can use any sort of tube, its just there to hold your cells as you centrifuge. The only factor you have to consider is if the tube you are using is able to be used at that speed in the centrifuge (some tubes aren’t strong enough for very high speeds).

  5. Lena on August 2, 2009 at 12:34 pm

    I got one transformant for one cell line when I plated out 50ul (and none for the other :(). I’m going to try and use them today anyway, and plate out all of the electroporated cells. One minor question about the protocol, what size rotor did you use?

    • Sean Smith on December 5, 2011 at 6:24 pm

      Hi Lena,

      I think the RCF may be pretty important. The protocol I used differs from this in a few respects and I only achieved 10^8 efficiency but I used 2000 g centrifugation speed. I don’t know, but perhaps too high-a-speed may damage the cells?

      Also, NEB recommends against adding additional 10% glycerol after removing the supernate in the final step. Simply resuspend in any remaining supernate which you could not remove and you’ll have the densest slurry (you’ll have less of it but it will be more efficient).

      Also, I used OD 600 = 0.5. You want the cells in their exponential phase of growth. This might well extend to 1.0 but you’d need to do another experiment to try to determine when growth begins to tail off for your cells/conditions. My cells were DH5a.

    • Sean Smith on December 5, 2011 at 6:28 pm

      Sorry, I should have said OD 600 = 0.5-0.7.

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