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Short answer
Transformation efficiency is the number of transformant colonies obtained per microgram of DNA, reported as colony-forming units per microgram (cfu/µg). It is calculated from a control plate: colonies counted, divided by the picograms of control plasmid (usually pUC19) in the transformation, multiplied by 106 pg/µg, and corrected for the fraction of the mixture you plated and any dilution.
Definition and units
Transformation efficiency describes how well a batch of competent cells takes up and maintains plasmid DNA. It is normalised to DNA mass so that different batches, protocols and suppliers can be compared. The unit is cfu/µg of DNA, and the standard test substrate is an intact, supercoiled control plasmid such as pUC19 (Origene protocol, Thermo Fisher E-Shot cuvettes manual, both read 2026-11-08).
Because a control plasmid is used, the figure is a property of the cells and the protocol under control conditions. It is not a prediction of how many colonies your own ligation will produce (see below).
Calculating transformation efficiency
Transformation efficiency equation
Two manufacturers publish the same calculation with slightly different terms. Origene (Suggested Transformation Protocol for competent cells, read 2026-11-08) gives:
Efficiency (cfu/µg) = (colonies ÷ pg pUC19 transformed) × (106 pg/µg) × (final volume of transformation mix, µl ÷ volume plated, µl)
Thermo Fisher (E-Shot cuvettes manual, read 2026-11-08) gives:
CFU/µg = (CFU on control plate ÷ pg pUC19) × (106 pg/µg) × (volume of transformants ÷ volume plated) × dilution factor
The Thermo form simply adds an explicit dilution factor for the case where the recovered cells were diluted before plating. With no dilution the factor is 1 and the two equations are identical.
Transformation efficiency calculation: two worked examples
| Term | Origene example | Thermo Fisher example |
|---|---|---|
| Colonies counted | 40 | 50 |
| pUC19 transformed | 20 pg | 5 pg |
| Volume of transformation mix | 1,000 µl (1 ml) | 500 µl |
| Volume plated | 5 µl | 100 µl of a 1:200 dilution |
| Dilution factor | not used | 2 × 102 |
| Calculation | 40 ÷ 20 × 106 × 1,000 ÷ 5 | 50 ÷ 5 × 106 × 500 ÷ 100 × 2 × 102 |
| Result | 4 × 108 cfu/µg | 1.0 × 1010 cfu/µg |
The arithmetic in both columns was re-checked for this page: 2 × 106 × 200 = 4 × 108, and 10 × 106 × 5 × 200 = 1 × 1010. The two examples differ by a factor of 25 because of the cells and method used (the Thermo example is an electroporation workflow), not because the formula differs.
The Origene protocol plates 5 µl of the final mixture on LB agar with 100 µg/ml ampicillin for the pUC19 control. Whatever your plating volume, use the same volume in the formula that you actually spread.
Transformation efficiency units
Express the result as cfu/µg. The 106 factor is only a unit conversion from picograms to micrograms, and the Origene and Thermo Fisher examples both enter the control plasmid in picograms.
Why the label number is not the number you will get
- Ligations are lower than control plasmid. Addgene (Bacterial Transformation protocol, read 2026-11-08) states that efficiencies are approximately 10-fold lower for ligation of inserts into vectors than for an intact control plasmid.
- Cell batches differ. Addgene notes that commercial competent cells range significantly in efficiency.
- DNA quality and volume matter. Thermo Fisher recommends DNA free of phenol, ethanol, salts, protein and detergents for maximum efficiency, and states that undiluted ligation mixtures or too high a DNA volume decrease efficiency (and raise the risk of arcing in electroporation). Addgene’s tip is that with 100-1000 ng of total DNA in a ligation, 1 µl of a 1:5 or 1:10 dilution often gives more colonies than 1 µl undiluted.
- Plasmid size. For large plasmids (over 10 kb) and BACs, Addgene says chemically competent cells are less efficient and suggests electrocompetent cells; see electroporation parameters and buffers.
The number on a vial is therefore a quality measure for the cells, and your own experiment should be compared against a positive control run in parallel.
No colonies after transformation: a checklist
For “why did I not get colonies after ligation transformation?”, the checks Addgene lists are the most direct, and they map back to the calculation above.
- Match the antibiotic. Confirm the plate contains the antibiotic for the resistance gene on your plasmid.
- Run a positive control. Transform a known intact plasmid in parallel. Colonies on the control plate with none on the experimental plate point toward the ligation or DNA (the cells and procedure worked). No colonies on the control point toward the cells or the procedure, and you can quantify the cells with the formula above.
- Reduce ligation input. Try a 1:5 or 1:10 dilution of the ligation rather than undiluted mix (Addgene); for electroporation, clean the DNA of salts first (Thermo Fisher recommends ethanol precipitation and resuspension in 0.5X TE).
- Allow outgrowth when the antibiotic needs it. Addgene says outgrowth can be skipped for ampicillin but should be at least 20-30 minutes for other antibiotics.
- Consider insert and vector size. Large constructs transform less efficiently in chemically competent cells.
Related methods pages: gene cloning, TA and TOPO cloning and bacterial transformation.
Griffith’s transforming principle
The word “transformation” comes from bacterial genetics. Avery, MacLeod and McCarty (J Exp Med 1944;79:137-158, read 2026-11-08) refer to Griffith’s original observation that heat-killed Type III pneumococci could cause rough (R) Type II variants to acquire the Type III capsule, and report that a purified fraction inducing this change “consists principally, if not solely, of a highly polymerized, viscous form of desoxyribonucleic acid”. The transforming principle was therefore identified as DNA, which is why the laboratory procedure of taking up plasmid DNA kept the name.
What are competent cells?
Competent cells are bacteria prepared so that they can take up foreign DNA, by chemical treatment or for electroporation. Efficiency is how you measure how good that preparation is. See the competent cells guide for preparation and handling.
Common misunderstandings
- A high cfu/µg on the vial does not guarantee a high colony count from a ligation (see the ten-fold figure above).
- More DNA is not always better; in highly competent cells less DNA can give higher efficiency (Addgene).
- Efficiencies from different formulas are comparable only if the volumes, dilution and plasmid mass are entered consistently.
FAQ
What is transformation efficiency measured in?
Colony-forming units per microgram of DNA (cfu/µg), usually determined with a control plasmid such as pUC19.
Why did I not get colonies after ligation transformation?
Check that the plate antibiotic matches the plasmid’s resistance gene, run a positive control, dilute the ligation, allow outgrowth if the antibiotic requires it, and note that ligations typically give about ten-fold fewer transformants than intact control plasmid (Addgene).
What was Frederick Griffith’s transforming principle?
Griffith’s observation was that a heat-killed virulent pneumococcal type could confer its capsule type on R variants. Avery, MacLeod and McCarty later showed the active fraction was principally DNA.
What are competent cells?
Bacterial cells prepared to take up DNA for transformation, by chemical or electroporation methods.
Sources
- Origene, Suggested Transformation Protocol (alpha-Gold competent cells), PDF, read 2026-11-08: cdn.origene.com/assets/documents/other/competentcells/transformationprotocolalphagold.pdf
- Thermo Fisher Scientific, E-Shot Electroporation Cuvettes manual, PDF, read 2026-11-08: documents.thermofisher.com/TFS-Assets/LSG/manuals/eshot_cuvettes_man.pdf
- Addgene, Bacterial Transformation protocol, read 2026-11-08: addgene.org/protocols/bacterial-transformation/
- Avery OT, MacLeod CM, McCarty M. J Exp Med 1944;79(2):137-158, PMC2135445, read 2026-11-08.
