Transformation efficiency

Transformation efficiency

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I accidentally spun down the cells after heat shock treatment before adding the media into the tubes while doing transformation? will the transformation work?

This is hard to answer since we don't know the exact conditions you chose, as these are rotation speed (or better g), how long you spun them down and also if you used a cooled centrifuge or not. It's also important how gentle you resuspended the pellet.

However, I would plate these cells, no matter what happened since you are at the last step of the procedure and have nothing to loose. Do the usual dilutions you use for plating your cells and then add another plate with undiluted transformation mix. If you have only very few surviving cells after your mistake you might get your clones on this plate. If it doesn't work, you will have to repeat the transformation.

The question is following: did you incubate cells on ice before spinning? Protocol, for example this one for $DH5alpha$, calls for just 10 minute incubation after heat shock:

Heat shock at exactly 42°C for exactly 30 seconds. Do not mix. Place on ice for 5 minutes. Do not mix. Pipette 950 µl of room temperature SOC into the mixture.

Did you centrifuge your tubes after step in bold? My bet is that spinning cells down (taking into account what @chris wrote about centrifugation parameters) will not dramatically decrease efficiency of transformation. DNA should be already in cells after step, highlighted in the quote.


Transformation efficiency should be determined under conditions of cell excess. Ώ] The number of viable cells in a preparation for a transformation reaction may range from 2×10 8 to 10 11 most common methods of E. coli preparation yield around 10 10 viable cells per reaction. The standard plasmids used for determination of transformation efficiency in Escherichia coli are pBR322 or other similarly-sized or smaller vectors, such as the pUC series of vectors. Different vectors however may be used to determine their transformation efficiency. 10–100 pg of DNA may be used for transformation, more DNA may be necessary for low-efficiency transformation (generally saturation level is reached at over 10 ng). ΐ]

After transformation, 1% and 10% of the cells are plated separately, the cells may be diluted in media as necessary for ease of plating. Further dilution may be used for high efficiency transformation.

Transformation efficiency can be measured in transformants or colony forming unit (cfu) per μg DNA used. A transformation efficiency of 1×10 8 cfu/μg for a small plasmid like pUC19 is roughly equivalent to 1 in 2000 molecules of the plasmid used being transformed. In E. coli, the theoretical limit of transformation efficiency for most commonly used plasmids would be over 1×10 11 cfu/μg. In practice the best achievable result may be around 2–4×10 10 cfu/μg for a small plasmid like pUC19, and considerably lower for large plasmids.

Transformation efficiency - Biology

Hi. I have a question about the transformation efficiency of competent E. coli. I know that the efficiency is calculated based on the amount of plasmid used in the transformation and the nuber of colones resulting. Does the initial volume of competent cells affect this at all? The maufacturer claims that using less than 100 ul of competent cells in my transformation procedure will reduce the efficiency of transformation. I don't understand this - if I also reduce the amount of plasmid I use, shouldn't the efficiency stay the same?

Does the efficiency calculation assume that every cell takes up a plasmid?

If my cells have an efficiency of < 10^9 cfu/ng can I just use more cells to start with and end up with a comperable level of colonies?
Thanks for your help.

The volume and the type of tube used both have effects on the transformation efficiency. This is because the final temperature the cells reach during the heat pulse is affected by these parameters. If the final temperature is too high, the cells die. If the final temperature is too low, the transformation efficiency is reduced. This was published in the early 80s in a BRL Focus article. I have successfully transformed cells with a plasmid containing a temperature-induced promoter construct by pulsing at 30C for 5 minutes. But I don't know if that time point was optimal.

The cells take up more than one plasmid and, in addition, the cell surface is coated with other copies of the plasmid (as well as excess copies of the insert DNA fragment since the ratio of insert to vector is usually greater than 1:1) that did not get taken up.

Invitrogen and Statagene have both published brochures on transformation efficiency and the number of colonies required to generate a cDNA library.

Calcium chloride protocols typically generate 5e6 to 2e7 cfu/ug of supercoiled plasmid DNA or 5000-10,000 cfu/ng of ligated DNA.
The Hanahan protocol generates 5e8 cfu/ug of supercoiled plasmid DNA or 500,000 cfu/ng of ligated DNA.
Electroporation protocols generate 1e9 to 1e10 cfu/ug of supercoiled plasmid DNA or 5,000,000-10,000,000 cfu/ng of ligated DNA.

There is a plasmid saturation level effect also. The cfu/ng plateau is surprisingly low, but will vary acording to the type of competency and the cell strain.

Thanks. If I understood your answer correctly, I don't have to worry too much about optimal efficiency if I am merely cloning for sequencing, rather than building libraries.

Manufacturer would be glad you to use 100 and more ul of cells! ))
I use 10 ul cells and 1 ul of plasmid for recloning perposes. Works always!

Transformation efficiency - Biology

By mixing E. coli cells with plasmid DNA carrying an antibiotic resistance gene, one can isolate cells that have incorporated the plasmid as an autonomously replicating DNA molecule. This is a naturally occuring process, called transformation, but it occurs at such a low frequency in nature that it is not feasible as a routine method for getting recombinant plasmids into E. coli . Much higher frequency of transformation can be obtained if E. coli cells are treated with a solution of divalent cations (usually Ca) at low temperatures to make them "competent" to take up plasmid DNA. It is thought that the Ca++ ions form an ionic bridge between the negatively charged phosphate groups of both the DNA and the membrane phospholipids. The cold temperature crystallizes the membrane stabilizing this interaction. When the cells are then exposed to a "heat shock" some of the cells take up the DNA. If the cells are allowed to warm up above 4 deg C before the heat shock step, they will not be competent. Be sure to use good sterile technique during the entire procedure.

Usually the number of cells in a transformation is in excess and the limiting "reagent" is the DNA. A measure of the quality of the competent cells is the transformation efficiency. This is the number of transformants (colonies which grow on the plate) obtained per ug of plasmid DNA. In order to determine the transformation efficiency, a control transformation with a specified amount of plasmid DNA is carried out along with the experimental transformation. The number of colonies is counted and multiplied by the appropriate dilution factor to get the total number of transformants. This is divided by the amount of DNA used in the transformation and expressed as transformants per microgram of DNA. Transformation efficiencies between 10^6 and 10^9 represent the normal range for competent E. coli cells.

E. coli cells are harvested by centrifugation when in late log phase of growth. They are chilled and treated with ice cold solutions containing relatively high concentrations (0.5 M) of divalent cations (usually Ca, but Mn also works well). To obtain the highest efficiency cells extraordinary care must be taken to keep cells cold during the preparation. The process involves multiple centrifugations and can by very tricky. Many researchers purchase aliquots of competent cells which have been prepared commercially and frozen at -80 C. You will be given an aliquot of such cells. The transformation efficiency of these cells should be approximately 10^9.

Procedure for Transforming E. coli Cells (Today we are using strain DH1-alpha cells purchased from Life Technologies)

1. Place 2 sterile 15 ml polypropylene tube on ice. Label one with your plasmid name and the other "control".

2. To each of the polypropylene tubes add 50 uL of the competent cells. Be sure to keep the cells cold during the transfer process.

3. Add 1 uL of your ligation mix to the first polypropylene tube and 1 uL of the control DNA (pUC18 at 10 pg/uL) to the control tube. Mix gently and incubate on ice for 30 minutes. Be sure to keep the cells cold during the transfer process.

4. Heat pulse the tubes in a 42 C water bath for 45 seconds exactly . The length of the heat pulse is critical for obtaining highest efficiencies.

5. Place tubes on ice for 2 min.

6. Add 0.9 mL of LB broth and incubate the tubes in a shaking incubator at 37 C for 60 minutes.

7. Label 3 LB/Amp plates with your plasmid name and 3 LB/Amp plates with "control". Number each set of plates 1-3.

8. Plate 0.1 mL of your plasmid transformation mix onto the LB/Amp plate labeled 1 and spread the broth out evenly over the plate. Also place 0.1 mL of your plasmid transformation mix into a second tube with 0.9 mL of LB broth. Mix and plate 0.1 mL of this mixture onto the LB/Amp plate labeled 2. Place 0.1 mL of the dilution from the second tube into a third tube with 0.9 mL of LB broth. Mix and plate 0.1 mL of this mixture onto the LB/Amp plate labeled 3.

9. Use the same plating procedure (step 8) to plate out the control transformation mix.

10. Incubate all plates at 37 C for 14-18 hours. Do not incubate any longer than 18 hours. After incubation is complete place plates in the refrigerator until needed.

Transformation efficiency - Help! (May/08/2006 )

I needed some big help in calculating transformation efficiency.

If the DNA concentration was 15 ng/ml and the plating volume was 100ul. The final volume at recovery was 100 ul. The number of transformants I got were 67. How can I calculate the transformation efficiency in units of transformants/ug of DNA.

I will really really appreciate any help in this.

Did you use 900ul of SOC or LB? Here is the example that I came across a while ago in some catalogue. Worh out like this

"After 100ul of competent cells are transformed with 0.1ng of uncut plasmid DNA, the transformation reaction is added to 900ul of SOC medium (0.1ng DNA/ml). A 1:10 dilution with SOC medium (0.01ng DNA/ml) is made, and 100ul is plated on each of two plates (0.001ng DNA/100ul). If 200 colonies are obtained (average of two plates), whats the transformation efficiency?

(200 cfu/0.001ng) x (1ng/ 10 to the power of -3 ug-microgram) = 2x10 to the power of 8 cfu/ug DNA

Sample 6a Transformation Lab

Bacterial transformation occurs when a bacterial cell takes up foreign DNA and incorporates it into its own DNA. This transformation usually occurs within plasmids, which are small circular DNA molecules separate from its chromosome. There can be 10 to 200 copies of the same plasmid within a cell. These plasmids may replicate when the chromosome does, or they may replicate independently. Each plasmid contains from 1,000 to 200,000 base pairs. Certain plasmids, called R plasmids, carry the gene for resistance to antibiotics such as ampicillin, which is used in this lab.

Plasmids function in transformation in two different ways. They can transfer genes that occur naturally within them, or they can act as vectors for introducing foreign DNA. Restriction enzymes can be used to cut foreign DNA and insert it into the plasmid vectors. The bacteria used in this lab were Escherichia coli (E. coli). It was ideal for this transformation study because it can be easily grown in Luria broth or on agar, and it has a relatively small genome of about five million base pairs.

Transformation is not the only method of DNA transfer within bacteria. Conjugation is a DNA transfer that occurs between two bacterial cells. A bridge is formed between the two cells and genetic information is traded. In transduction, a virus is used to transfer foreign DNA into a bacterial cell.

The transformed E. coli with the ampicillin resistance gene will be able to grow in the ampicillin plates, but the non-transformed E. coli will not.

The materials needed for this lab were 2 sterile test tubes, 500 μL of ice cold 0.05M CaCl2, E. coli bacteria cultures, a sterile inoculating loop, a sterile micropipette, 10 μL of pAMP solution, a timer, ice, a water bath, 500 μL of Luria broth, a spreading rod, 4 plates: 2 ampicillin+ and 2 ampicillin – , and an incubator.

One sterile tube was labeled “+” and the other “-“. A sterile micropipette was used to transfer 250 μL of ice cold 0.05M CaCl2 to each tube. A large colony of E. coli was transferred with an inoculating loop to each tube. The suspension was then mixed by repeatedly drawing and emptying a sterile micropipette. 10μL of pAMP solution was added to the cell suspension in the tube marked “+” and mixed by tapping the tube. Both tubes were immediately put on ice for 15 minutes and then soaked in a 42° C water bath for 90 seconds. The tubes were then returned to ice for another 2 minutes.

After the heat shock, 250 μL of Luria broth were added to each tube. The tubes were mixed by tapping. Two plates of ampicillin + agar were labeled LB/AMP+ and LB/AMP-. The two plates of ampicillin- agar were labeled LB+ and LB-. 100 μL of the cell suspension in the “+” tube were placed on the LB+ and the LB/AMP+ plates. 100μL of the cell suspension in the “-” tube were added to the LB- and the LB/AMP- plates. These were spread with a spreading rod that was sterilized by passing it over a flame after each use. The plates were allowed to sit for several minutes and then incubated over night inverted at 37° C.

1. Compare and contrast the number of colonies on each of the following pairs of plates. What does each pair of results tell you about the experiment?
LB+ and LB- Both of these plates had a lawn of bacteria. This proves that the bacteria are capable of growing on the agar and that there was nothing preventing growth beside the ampicillin.

LB/AMP- and LB/AMP+ The LB/AMP- had no growth, but the LB/AMP+ had small growth. This shows that the bacteria was transformed and developed a resistance to ampicillin.

LB/AMP+ and LB+ The LB/AMP+ had less growth than the LB+. This shows that the transformation was not completely effective and only transformed some of the most competent bacterial cells.

2. Total mass of pAMP used = 0.05 μg

Total volume of cell suspension = 510 μL

Fraction of cell suspension spread on the plates = 0.196

Mass of pAMP in cell suspension = 0.0098

Number of colonies per μg of plasmid = 0.0294

3. What factors might influence the transformation efficiency? Explain the effect of each you mention.
Transformation efficiency could be affected by the size of the colony added to the solution. In a larger colony the efficiency would increase because there would be more receptive cells. Another factor would b the amount of pAMP added. The more pAMP added, the higher the efficiency. The amount of Luria broth added could also affect efficiency. If the amount of Luria broth was increased, the efficiency would decrease.

Error Analysis:
This lab had several steps, each giving the potential for error. All of the measurements had to be precise and accurate, and the heat shock timing was also a very complicated procedure. Error in this lab could have been caused by the concentration of the CaCl2 due to the fact that most of it was frozen.

Discussion and Conclusion:
The bacteria treated with the pAMP solution developed a resistance to ampicillin and were able to grow on the ampicillin+ plate. Those that were not treated with the pAMP were not able to grow on this medium. The plates with no ampicillin served as a control to show how the bacteria would look in normal conditions. Transformation is never fully effective, Only cells that are competent enough are able to take up the foreign DNA. Therefore, the ampicillin + plates showed less growth than the control plate.

Transformation Efficiency - Factors Affecting Transformation Efficiency

A number of factors may affect the transformation efficiency:

Plasmid size — A study done in E. coli found that transformation efficiency declines linearly with increasing plasmid size. Individual cells are capable of taking up many DNA molecules, and that the presence of multiple plasmids does not significantly affect the occurrence of successful transformation events.

Forms of DNA — Supercoiled plasmid have a slightly better transformation efficiency than relaxed plasmids which are transformed at around 75% efficiency of supercoiled ones. Linear and single-stranded DNA however have much lower transformation efficiency. Single-stranded DNAs are transformed at 104 lower efficiency than double-stranded ones.

Genotype of cells — Cloning strains may contain mutations that improve the transformation efficiency of the cells. For example, E. coli K12 strains with the deoR mutation, originally found to confer an ability of cell to grow in minimum media using inosine as the sole carbon source, have 4-5 times the transformation efficiency of similar strains without. For linear DNA, which is poorly transformed in E. coli, recBC or recD mutation can significant improve the efficiency of its transformation.

Growth of cellsE. coli cells are more susceptible to be made competent at a particular stage of their growth cycle, possibly when the cell volume is the greatest. When preparing competent cells, cells are therefore harvested at particular optical density (normally around 0.4, higher value of 0.94-0.95 may be used but impractical when cell growth is rapid.)

Methods of transformation — The method of preparation of competent cells, the length of time of heat shock, temperature of heat shock, and various additives, all can affect the transformation efficiency of the cells. The presence of contaminants as well as ligase in a ligation mixture can reduce the transformation efficiency, and heat inactivation of ligase may be necessary for electroporation. Normal preparation of compentent cells can yield transformation efficiency ranging from 106 to 108 cfu/μg DNA. Protocols for chemical method however exist for making supercompetent cells that may yield a transformation efficiency of over 1 x 109. Electroporation method in general has better transformation efficiency than chemical methods with over 1 x 1010 cfu/μg DNA possible, and it allows large plasmids of 200 kb in size to be transformed.

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Collection of immature embryos that are healthy and at the correct developmental stage is a key factor for wheat transformation. The conditions of the controlled environment growth rooms should be adjusted to grow plant material vigorously so that the immature embryos reach the early milk stage, GS73 (Zadoks, Chang, & Konzak, 1974 ), ∼14 days post anthesis. In addition, particular care needs to be taken to reduce the risk of spreading pathogens. The parent plants should not be sprayed with insecticides or fungicides at any stage of growth, and therefore high levels of plant hygiene must be maintained. As the plant material develops to the right developmental stage over 12-14 weeks, the seeds from the variety to be transformed should be sown every week in small batches. This will ensure a continuous supply of immature embryos at the right developmental stage for transformation.


This basic protocol describes growing of donor plants used to produce immature embryos used in the transformation system.

NOTE: Sequentially sow small batches of wheat seeds at weekly intervals to ensure a continued supply of immature embryos. Donor plants are grown at 20 ± 1°C day and 15 ± 1°C night temperatures, 70% humidity, with light levels of 600 μmol m −2 s −1 provided by fluorescent tubes and tungsten lighting in controlled environment growth rooms.


  • Controlled environment growth room
  • 24CT seed tray inserts with 5-cm-square cells
  • Seed tray
  • Small dibber (or pencil)
  • Medium dibber (optional)
  • 13-cm-diameter pots
  • Watering can fitted with a fine rose

1. Place a 24CT insert inside a seed tray and fill with John Innes cereal mix.

2. Using a small dibber (or pencil) make a hole ∼1.5-2 cm deep in the center of the cereal mix within each module

3. Place a single wheat seed in each hole and cover with a little more cereal mix (if needed). Tap the tray to level the cereal mix.

4. Using a watering can with a fine rose fitted, gently water the seeds.

5. Place the tray in a controlled environment room set to the conditions listed above (note). Plants should be monitored daily. The compost should be kept moist but not waterlogged.

Seedlings will be visible within a week of sowing. After ∼30 days, the seedlings will be large enough to transplant.

6. Filling 13-cm-diameter pots with John Innes cereal mix.

7. Make a small hole in the center of the compost using a medium-sized dibber or your finger. The hole should be about the size of the 5-cm modules in which your wheat seedlings germinated.

8. Water the seedlings that are about to be transplanted. Loosen the seedling roots from the module's walls by gently squeezing and release the seedling.

9. Keep the root ball intact as much as possible by supporting the seedling from underneath. Gently plant the seedling into the hole in the fresh compost. Firm the compost around the roots gently.

10. Water using a watering can with a fine rose fitted to it.

11. Grow the plants in a controlled environmental room under the conditions listed above until they produce donor embryos (see Basic Protocol 3 and Time Considerations). Continue to monitor plants daily. The compost should be kept moist but do not overwater.


We recommend using a reporter-gene construct in your initial experiments for optimization of the protocol to your local conditions. The pGoldenGreenGate (pGGG) we describe here contains the hygromycin resistance gene (hpt) and Cat1 intron under the control of a rice Actin1 promoter, and the β-glucuronidase gene (GUS) with two introns (GUS2Int) under the control of the rice ubiquitin promoter (Addgene cat. no. 165418), along with the helper plasmid pAL155 containing an additional VirG gene (Fig. 1). This combination gave the highest transformation efficiency within our system (Hayta et al., 2019 ).

NOTE: The following steps should be carried out in a laminar flow cabinet under aseptic conditions. Ensure cells, cuvette, and DNA are chilled on ice.


  • Electrocompetent cells of hypervirulent Agrobacterium tumefaciens strain AGL1
  • Binary vector DNA to be used
  • Helper plasmid DNA, if using
  • LB medium, solid and liquid (Bertani, 1951 )
  • Antibiotics to select for the Agrobacterium strain and binary vector used (e.g., rifampicin or kanamycin see step 7)
  • MG/L medium, liquid
  • Wheat inoculation medium (WIM), liquid
  • 100 mM acetosyringone (AS see recipe)
  • Electroporation cuvette with 2-mm separation (Sarstedt Limited 67.742), prechilled on ice
  • Bio-Rad GenePulser or similar electroporation device
  • 15-ml Falcon tubes (Life Sciences 430790), sterile
  • Parafilm strips (Parafilm®-PM-992)
  • Electric shaker
  • 90-mm single-vent petri plates (Thermo Scientific 101R20)
  • 50-ml Falcon tube (Starlab E1450-0800)
  • Aluminum foil

Transformation of Agrobacterium with vector

1. Thaw 50 µl electrocompetent cells of hypervirulent Agrobacterium tumefaciens strain AGL1 on ice. This should take ∼5 min.

2. Add 1 µl (∼100 ng) binary vector DNA (and the same amount of helper plasmid, if using) to the competent cells, gently mix by flicking, and place back on ice.

3. Transfer the cells and DNA mixture to a prechilled cuvette on ice. Gently tap to ensure the contents are at the bottom of the cuvette. Maintain on ice.

4. Electroporate using a Bio-Rad GenePulser, with the settings 2.50 kV, 25 μFD, 400 Ω, and time constant between 8.0 and 9.5 ms.

5. Immediately add 100 µl liquid LB medium to the cuvette to allow cells to recover. Transfer the cells to 500 µl LB medium in a sterile Falcon tube. Wrap the lid of the Falcon tube with Parafilm to secure.

6. Incubate the cells at room temperature, lay the Falcon tube horizontally on a shaker, and gently shake at 100 rpm for 4-6 hr.

7. Spread 20 and 40 µl of the electroporation culture, respectively, onto two plates solid LB medium supplemented with appropriate antibiotics.

For C58 strains of Agrobacterium (such as AGL1), add rifamycin at 50 µg/ml for selection, and when using pGGG, use kanamycin at 50 µg/ml.

8. Incubate the plate upturned at 28°C for ∼48 hr.

Preparation of Agrobacterium for transformation

9. From the inoculated plate, select single colonies of Agrobacterium AGL1, containing the desired vector, and inoculate 10 ml liquid LB medium containing the appropriate antibiotics. Incubate at 28°C in a rotary shaker, shaken at 200 rpm for ∼65 hr.

10. Prepare Agrobacterium standard inoculums for transformation as previously described by Tingay et al. ( 1997 ). Mix equal quantities of the Agrobacterium culture and 30% sterile glycerol. Make 400-µl aliquots in 0.5-ml microcentrifuge and freeze at −80°C.

The standard inoculum aliquots are stored at −80°C until required. They can be stored almost indefinitely.

11. The day before wheat transformation, use a single 400-µl standard Agrobacterium inoculum to inoculate 10 ml of MG/L (Garfinkel & Nester, 1980 ) liquid medium without antibiotics and incubate at 28°C in a rotary shaker, shaken at 200 rpm overnight (∼16 hr).

12. On the day of transformation, pellet the bacteria by centrifugation in a 50-ml Falcon tube at 3100 rpm for 10 min at 24°C. Discard the supernatant, and resuspend the cells gently in 10 ml wheat inoculation medium (WIM) to an optical density of 0.5 OD (600 nm). Then, add 100 mM acetosyringone (AS) to 100 µM final concentration.

13. Incubate the culture for 4-6 hr at room temperature with gentle agitation (80 rpm) in the dark (wrap Falcon tube with aluminum foil) before proceeding to Basic Protocol 3.


This protocol describes the collection of starting material at the correct stage, their sterilization, and then the inoculation of immature embryos.


  • Donor wheat plants with spikes (Basic Protocol 1)
  • 70% (v/v) ethanol
  • Distilled water, sterile
  • 10% (v/v) sodium hypochlorite (Fluka 71696)
  • Wheat inoculation medium (WIM see recipe)
  • Silwet L-77
  • Wheat co-cultivation medium plates (see recipe)
  • 150-ml Sterilin jar
  • Fine-point forceps
  • 1.7-ml microcentrifuge tubes
  • 90-mm-diameter single-vent petri plates (Thermo Scientific No 101R20)
  • 50-mm-diameter petri plate, sterile
  • Scissors
  • Micropore tape

Collection and sterilization of immature seeds

1. Collect the wheat spikes ∼14 days post anthesis, when the immature embryos are 1-1.5 mm in diameter and at the early milk stage GS73 (Fig. 2A).

2. Use kernels from florets 1 and 2 (Fig. 2B) on central spikelet (Fig. 2C) for transformation. Cut off the awns from the ears ∼3-5 mm from the grain.

The seed coat can be removed, but this is not essential unless contamination problems are expected.

3. Separate the immature grains from the ear and place in a 150-ml Sterilin jar.

4. Sterilize the grains under aseptic conditions (e.g., laminar flow cabinet) using 70% (v/v) ethanol for 1 min. Rinse once with sterile distilled water and then place in 10 ml 10% (v/v) sodium hypochlorite and let stand 7 min. Then wash the grains three times with sterile distilled water.

Isolation of immature embryos, inoculation with Agrobacterium, and co-cultivation

5. Isolate the embryos (Fig. 3A) from the immature grains using fine forceps.

6. Transfer embryos to 1.7-ml microcentrifuge tubes containing 1 ml WIM with 0.05% Silwet L-77, placing ∼100 embryos in each tube (Fig. 3B).

7. After isolating all the embryos, remove the WIM and add fresh WIM to the microcentrifuge tube(s). Centrifuge the isolated embryos 10 min at 14,000 rpm, 4°C (Ishida, Tsunashima, Hiei, & Komari, 2015 ).

8. Remove WIM with a pipet, add 1 ml Agrobacterium solution, and invert the tubes repeatedly for 30 s. Incubate at room temperature for at least 20 min.

9. After the incubation period, pour the Agrobacterium suspension with the embryos into an empty sterile 50-mm-diameter petri plate and then remove the suspension with a pipet.

10. Transfer 25 embryos, scutellum side up, to a fresh plate(s) of wheat co-cultivation medium in 90-mm-diameter single-vent petri plates.

11. Seal the petri plates with Micropore tape and incubated at 24 ± 1°C in the dark for 3 days of co-cultivation (Fig. 3C).


This protocol describes the resting period immediately after transformation, the selection process, plantlet regeneration and rooting, and finally acclimation from in vitro culture to soil. The resting period is a 5-day period following co-cultivation in which the Agrobacterium is removed by using timentin however, the plant material is not subjected to selection pressure. This resting period allows the material to recover from the Agrobacterium infection and initiates callus induction. The selection process that follows puts the calli produced from the immature embryos under selection pressure using the antibiotic hygromycin. Cells that contain the T-DNA and are therefore resistant to hygromycin can proliferate on medium containing hygromycin. The selection pressure continues through the remaining in vitro steps however, it is these first two phases that are referred to as Selection 1 and 2. During these two selection stages it is important that subcultures are done at the scheduled time, as delay may result in the growth of latent Agrobacterium, causing overgrowth and browning of callus, and/or the reduced stringency of selection may result in the growth of “escapes”, i.e., non-transformed plants.


  • Inoculated wheat embryos (Basic Protocol 3)
  • Wheat callus induction medium (WCI see recipe), solid and liquid
  • Wheat regeneration medium (WRM see recipe), solid
  • Extract-N-Amp™ Plant Tissue PCR Kits (Sigma XNAP-1KT)
  • REDExtract-N-Amp PCR Reaction Mix (Sigma XNAS)
  • GUS solution (see recipe)
  • 90-mm-diameter petri plate (Thermo Scientific No 101R20)
  • Deep petri dishes (tissue culture dish, 90 mm diameter × 20 mm, Falcon 353003)
  • De Wit culture tubes (Duchefa W 1607)
  • De Wit tray (Duchefa T1608)
  • Clear plastic propagator lid
  • Micropore tape

Resting period, callus induction, and selection of transformed material

1. After 3 days of co-cultivation, excise the embryogenic axes from the embryos using forceps (Fig. 4A).

2. Transfer the embryos to fresh wheat callus induction medium plates (WCI) and incubate at 24 ± 1°C in the dark for 5 days.

Timentin is included in the WCI medium to control Agrobacterium growth during the resting period.

3. Selection 1: After the 5-day resting period, transfer the embryos, scutellum side up, to fresh WCI plates as described above with the addition of 15 mg/liter hygromycin (10 mg/liter for Cadenza Fig. 4B), and incubate at 24 ± 1°C in the dark for 2 weeks (Fig. 4C).

4. Selection 2: After 2 weeks, split the calli at the into clumps of ∼4 mm 2 . Callus pieces derived from a single embryo should be labelled to keep track of their origin. Transfer the calli to fresh selection plates (WCI plates) as above, but with 30 mg/liter hygromycin (15 mg/liter for Cadenza), reducing the number of calli per plate by approximately half. Incubate at 24 ± 1°C in the dark for 2 weeks (Fig. 4D).

5. After 2 weeks, transfer the calli to a lit culture room under fluorescent lights (100 μmol/m 2 /s) at 24 ± 1°C with a 16-hr photoperiod and covered with a single layer of paper towel for a further week.

During this period, putative transformed lines should start to green and produce small shoots (Fig. 5A).

The hygromycin level might vary depending on the variety being transformed (see medium recipes). We recommend making a kill curve when trying to set up a completely new transformation system with varieties other than those listed here.

Regeneration of transgenic plants

6. After the 3 weeks on the selection 2 plates, transfer the calli a final time to wheat regeneration medium (WRM) in deep petri dishes (90 mm diameter × 20 mm). Remove the tissue paper covering the plates. Culture calli under fluorescent lights (100 μmol/m 2 /s) at 24 ± 1°C with a 16-hr photoperiod.

When transforming different wheat varieties, Kronos and Cadenza prefer slightly different regeneration media (see medium recipes below). Also, the concentration of hygromycin is reduced to 15 mg/liter for Cadenza.

If shoots/calli need to grow for longer on WRM, culture should be subcultured onto fresh medium.

7. Transfer regenerated shoots that are 1-2 cm in length with visible roots (Fig. 5B) to De Wit culture tubes containing 8 ml WCI without growth regulators and supplemented with 15 mg/liter hygromycin.

Label all plantlets derived from single embryos to keep track of their origins.

Putative transformed plants develop a strong root system with root hairs (Fig. 5C).


8. After ∼10 days, gently remove regenerated plantlets with strong root systems from the tubes using long forceps. Wash the roots with cool running water to remove any remaining tissue culture medium.

9. Plant the plants in cereal mix in 24CT trays and cover with a clear plastic propagator lid. The plants should remain covered with the propagator lid for ∼1 week to maintain high humidity around them while they become established in soil.

The plants are grown under the same conditions as donor plants within a controlled environment room.

10. Once the plants are well established in the soil, leaf samples can be collected for further analysis to confirm the presence of the introduced genes.


Successful transformation should be confirmed by PCR amplification of the hygromycin resistance gene and/or copy number analysis performed by quantitative PCR.

DNA extraction

11. Harvest 0.5- to 0.7-cm leaf samples into PCR tubes, and extract DNA with Extract-N-Amp™ Plant Tissue PCR Kits following the manufacturer's instructions.

HygR (hpt) polymerase chain reaction (PCR)

12. Amplify the 335-bp amplicon of the hygromycin resistance gene (hpt) by PCR using the primer pair HygF 5′-AGGCTCTCGATGAGCTGATGCTTT-3′, HygR 5′-AGCTGCATCATCGAAATTGCCGTC-3′, along with REDExtract-N-Amp PCR Reaction Mix (cat. no. XNAS), in a 20-µl volume per reaction. Each reaction should comprise 10 µl 2× PCR Reaction Mix (REDExtract-N-Amp), 4 µl DNA extract (∼50 ng DNA), 1 µl (10 mM) of each primer (Hyg F and Hyg R), and 4 µl sterile laboratory-grade water, in 20 µl total volume.

13. Perform PCR in a thermal cycler under the following conditions:

Initial step: 3 min 95°C
34 cycles: 30 s 95°C (denaturation)
30 s 58°C (annealing)
1 min 72°C (extension)
Final extension: 7 min 72°C
Final hold: - 10°C.

14. Resolve the PCR products by gel electrophoresis on a 1% agarose gel containing 0.1 μg/ml ethidium bromide.

GUS histochemical assay

NOTE: We suggest carrying out initial experiments with a control vector expressing a reporter gene such as GUS to assess and optimize the system for your local conditions. Using a reporter construct will allow you to follow the transformation process through the different stages.

15. Immerse the plant material in GUS assay substrate at 37°C under dark conditions overnight (∼ 16 hr).

16. Remove the GUS solution, and for green plant material, immerse in 70% ethanol to remove chlorophyll. Leave all green samples in 70% ethanol to remove chlorophyll and other plant pigments prior to visualization and photography.

Determine GUS activity after co-cultivation, resting (Fig. 6A), Selection 1 (Fig. 6B), and rooting in medium (Fig. 6C). An example of transgene inheritance in T1 seeds in the next generation as demonstrated by a GUS histochemical assay is shown (Fig. 6D).

Improved in planta genetic transformation efficiency in bitter gourd (Momordica charantia L.)

Production of transformed bitter gourd plants through in vitro regeneration is a laborious practice, which may also result in somaclonal variations. Thereby, in the current study, we have established a less tedious in planta protocol for more efficient genetic transformation of bitter gourd var. NBGH-470 (Apurva) using the seed as a target material. Bitter gourd seeds were transformed by Agrobacterium tumefaciens strain LBA4404 bearing the pCAMBIA1301 binary vector, and the transformed plants were selected against hygromycin B. The putatively transformed bitter gourd (To) plants were examined by GUS assay. Two parameters, including vacuum infiltration and sonication improving the in planta transformation, have been investigated in the present work. Amongst several time durations examined, the highest transformation rate (37%) was achieved upon subjecting the pre-cultured bitter gourd seeds to sonication for 15 min, followed by vacuum infiltration for 6 min in LBA4404 culture suspension. The transformed bitter gourd (T1) plants were selected against hygromycin B, and the transgene integration was evaluated by polymerase chain reaction (PCR), Southern hybridization, and reverse transcriptase PCR (RT-PCR). The developed protocol in the current study is suitable to genetically engineer the bitter gourd plants with disease and pest-resistant traits.

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Transformation process allows a bacterium to take up genes from its surrounding environment that is transformation involves the direct uptakes of fragments of DNA by a recipient cell and the acquisition of new genetic characteristics. There are two major parameters involved in efficiently transforming a bacterial organism. The first is the method used to induce competence for transformation. The second major parameter is the genetic constitution of the host strain of the organism being transformed. Competent cells are capable of uptaking DNA from their environment and expressing DNA as functional proteins. If a bacterium is said to be competent, it has to maintain a physiological state in which it can take up the donor DNA. Calcium chloride treatment is one of the best methods for the preparation of competent cells. Competence results from alterations in the cell wall that makes it permeable to large DNA molecules. This is a naturally occurring process and through this bacteria can transfer advantageous characteristics, such as antibiotic resistance. Bacteria can take DNA from the environment in the form of plasmid. Most of them are double stranded circular DNA molecules and many can exist at very high copy numbers within a single bacterial cell. Many naturally occurring plasmids carry an antibiotic resistant gene referred to as a marker.

In the process of transformation, the competent cells are incubated with DNA in ice. Then it is placed in a water bath at 42ºC and further plunging them in ice. This process will take up the DNA into the bacterial cell. Then it is plated in an agar plate containing appropriate antibiotic. The presence of an antibiotic marker on the plasmid allows for rapid screening of successful transformants. Blue &ndashwhite selection (Alpha complementation) can be used to determine which plasmids carry an inserted fragment of DNA and which do not. These plasmids contain an additional gene (lac Z) that encodes for a portion of the enzyme &beta &ndash galactosidase. When it transformed into an appropriate host, one containing the gene for the remaining portion of &beta &ndashgalactosidase, the intact enzyme can be produced and these bacteria form blue colonies in the presence of X &ndash gal (5-bromo-4-chloro-3-indoyl-b-D-galactoside) and a gratuitous inducer called IPTG (Isopropyl &beta-D- Thiogalactopyronoside). These plasmids contains a number of cloning sites within the lac Z gene, and any insertion of foreign DNA into this region results in the loss of the ability to form active &beta &ndashgalactosidase. Therefore colonies that carry the plasmid with the insert, ie, Transformants will remain white and the colonies without the foreign DNA (Non-Transformants) will remain Blue. We can also calculate the efficiency of transformation by using the concentration of DNA and number of transformed colonies.

Watch the video: How to Calculate Transformation Efficiency (February 2023).