Scientist with gloved hands is scraping a petri dish with bacterial colonies

Bacterial transformation is a primary technique in molecular cloning to produce multiple copies of a recombinant DNA molecule. Prior to transformation, recombinant plasmids are constructed by inserting the DNA sequence of interest into a vector as described in traditional cloning basics. In transformation, the DNA, typically in the form of a plasmid, is introduced into a competent strain of bacteria so that the bacteria may then replicate the sequence of interest in amounts suitable for further analysis and/or manipulation.

The four key steps in bacterial transformation are

Infographic illustrates the four main bacterial transformation steps: 1) competent cell preparation, 2) transformation of cells, 3) cell recovery, and 4) cell plating
Figure 1. Bacterial transformation steps. 1) Competent cell preparation, 2) transformation of cells, 3) cell recovery, and 4) cell plating.

Competent cell preparation

E. coli is the most common bacterial species used in the transformation step of a cloning workflow. In its natural state, the competency of E. coli is very low (10-5−10-10) [1] thus cells must be made competent for efficient transformation. The protocols for preparing competent cells vary by choice of transformation method—heat shock or electroporation. Heat shock transformation is also known as chemical transformation and calcium chloride transformation. In both cases, a starter culture is prepared by inoculating a single fresh colony from an agar plate into a liquid medium (Figure 2). This starter culture and the subsequent sub-cultures are carefully monitored for active growth by continually measuring optical density at 600 nm (OD600). To obtain high transformation efficiency, it is crucial that cell growth is in mid-log phase at the time of harvest—which generally occurs at OD600 between 0.4 and 0.9; the optimal value depends on the culture volume, strain, and protocol. Harvested cells are then processed for heat shock treatment or electroporation (Figure 2).

Heat shock transformation: The harvested cells are incubated in calcium chloride (CaCl2) to make the cell membrane more permeable [2,3]. To further improve competency, Ca2+ ions may be supplemented or substituted with other cations and reagents, such as manganese (Mn2+), potassium (K+), cobalt ([Co(NH₃)₆]3+), rubidium (Rb+), dimethyl sulfoxide (DMSO), and dithiothreitol (DTT) [4].

Electroporation: The harvested cells are repeatedly washed with ice-cold deionized water by pelleting and resuspension to remove salts and other potential interfering components. After 3–4 wash cycles, the cells are pelleted and resuspended in 10% glycerol for storage [5].

Diagram of competent cell preparation for chemical transformation and electroporation: 1) cell growth, 2) cell preparation, and 3) aliquoting and storage

Figure 2. Preparation of chemically competent and electrocompetent cells.

Once prepared, competent cells should be evaluated for transformation efficiency. The transformation efficiency of competent cells is usually measured by the uptake of sub saturating amounts of a supercoiled intact plasmid, approximately 10–500 pg of pUC DNA. The results are expressed as the number of colonies formed (transformants), or colony forming units (CFU), per microgram of plasmid DNA used (CFU/μg) (see Cell Plating). The prepared cells can be stored for future use.

Plasmid transformation

The term “plasmid” was first used by Joshua Lederberg in his 1952 publication and adopted by scientific community to describe a genetic element that is not an organelle, virus, or other cell genetic element. The first plasmid was edited in 1973 demonstrating ability to replace one antibiotic resistant gene (tetracycline) to another (kanamycin) [6]. But the first true vector—plasmid that was used for delivering and manipulating foreign DNA inside a host cell—that became an essential tool in molecular biology was pBR322. Therefore, foreign DNA transfer aided by vectors is called plasmid transformation.

Plasmid uptake by chemically competent cells is facilitated by heat shock, and plasmid uptake by electrocompetent cells is facilitated by electroporation. The choice of method depends on the application, desired transformation efficiency, experimental goals, and available resources; refer to the  Competent Cell Selection Guide. When ready for the transformation step, competent cells should be thawed on ice and handled gently to retain viability. Cells can be mixed by gentle shaking, tapping, or pipetting, but vortexing should be avoided. 

Chemical transformation

When cations, such as Mg2+, are used to introduce plasmids into chemically competent cells, the process is called chemical transformation. This video demonstrates how to perform chemical transformation.

In chemical transformation, plasmid DNA is mixed with chemically competent cells, then briefly exposed to an elevated temperature—a process known as heat shock (Figure 3A). The initial incubation of cells and plasmid is carried out in a polypropylene tube on ice for durations ranging from 5–30 minutes.

For successful chemical transformation, 50–100 µL of competent cells and 1–10 ng of DNA is recommended. When a ligation mixture is used as the source for transforming DNA (often 1–5 µL is sufficient), purification prior to chemical transformation is generally not required. However, ligation mixtures may result in transformation efficiencies as low as 1–10%, compared to transformation with a supercoiled intact plasmid DNA.

Heat shock is performed at 42°C for 30 seconds as appropriate for the bacterial strain and DNA used. For smaller volumes of cells in smaller tubes, the heat shock interval, which depends on the surface-to-volume ratio of the cell suspension, should be reduced. Heat shocked cells are then returned to ice for ≥2 minutes before the next step (Figure 3A).

Diagram of chemical transformation and electroporation

Figure 3. Bacterial transformation using (A) chemically competent cells and heat shock, and (B) electrocompetent cells and electroporation.

Electroporation transformation

Bacterial transformation aided by electroporation is called electroporation transformation; electroporation involves using an electroporator to subject competent cells and the plasmid carrying DNA construct to a brief pulse of a high-voltage electric field (Figure 3B). This treatment induces transient pores in cell membranes, which permits plasmid entry into the cells (Figure 4). The most common type of electric pulse in bacterial transformation is exponential decay, where a set voltage is applied and allowed to decay over a few milliseconds, called the time constant(Figure 4A). The applied voltage is determined by field strength (V/cm), where V is the initial peak voltage and cm is the measurement of the gap between the electrodes of the cuvette used. Typically, electroporation of bacteria utilizes 0.1 cm cuvettes (20–80 µL volume) and requires a field strength of >15 kV/cm.

Panel A is a graph showing the decay of electric pulse with respect to time and panel B is a diagram of the four steps of the electroporation process for bacterial transformation

Figure 4. Bacterial transformation via electroporation. (A) Graph demonstrating exponential decay of electric pulse over time. (B) Workflow steps of the electroporation process.

One of the main issues with electroporation is arcing, or electric discharge, which may lower cell viability and transformation efficiency. Arcing often results from electroporation in conductive buffers, such as those containing MgCl2 and phosphates.

After transformation, unused competent cells may be refrozen. However, this will lower transformation efficiencies by about 50% for each freeze/thaw cycle. For best results, aliquot the cells after initial preparation into single-use volumes to minimize freezing and thawing. A single-use format is commercially available to enable transformation and recovery in the same tube and to circumvent the need for freezing and thawing of the cells. To refreeze unused cells, quickly freeze them in a dry ice/ethanol bath for 5 minutes, and store at –70°C. Avoid freezing or storing the cells in liquid nitrogen, which drastically reduces viability.

Cell recovery

Following heat shock or electroporation, transformed cells are cultured in antibiotic-free liquid medium for a brief period. This allows the expression of antibiotic resistance gene(s) from the introduced plasmid (Figure 5). This step improves cell viability and cloning efficiency. For electroporated cells, it is recommended that the cells are grown as soon as possible, since electroporation buffers are not formulated for long-term cell survival.

In the recovery step, prewarmed SOC media is added on the top of transformed cells (250 µL to 1 mL depending on protocol) and cultured at 37°C by shaking (225 rpm) for 1 hour. SOC medium, which contains glucose and MgCl2, is recommended to maximize transformation efficiency [4]. Use of SOC medium, instead of Lennox L Broth (LB Broth), has been shown to increase 2- to 3-fold formation of transformed colonies [7]. Strains for propagating bacteriophage M13 vectors do not require this step.

Diagram of cell growth and plasmid propagation in recovery step

Figure 5. Cell growth during the recovery step.

Cell plating

After growing in SOC medium, the cells are plated on solid media, typically LB agar plates with antibiotic(s) or other agents that serve as markers to identify and recover successful transformants. For example, if blue-white screening is performed, X-Gal and IPTG must be included in the agar plate. To ensure that the antibiotic is active, avoid using agar plates more than a few weeks old or days in some cases. Before cell plating, the plates should be prewarmed to a favorable growth temperature and be free of condensation to prevent contamination and mixed colonies.

The number of cells plated should produce enough individual, distinct colonies for further screening, avoiding overly crowded colonies. Cells cultured in SOC medium may be pelleted by centrifugation for 5 minutes at 600–800 x g and resuspended in a smaller volume for plating. For a 100 mm plate, a cell suspension of 100–200 µL is appropriate. If very few or very high number of colonies are present on the agar plate, please follow-up with troubleshooting recommendations.

Transformant recovery and clone selection

The culture plates are examined the next day for colony formation. Prolonged incubation should be avoided, as it often results in fusion of large colonies and the appearance of smaller, antibiotic-sensitive surrounding colonies, called satellite colonies, due to antibiotic breakdown around large colonies.

Transformed E. coli colonies then need to be isolated and expanded to isolate and analyze individual constructs. Depending on experimental goal, several or several dozen colonies would need to be picked up and expanded. Usually, colonies are picked up by pipette tip and inoculate in <5 mL LB or TB medium with appropriate selective antibiotic for growth to use for small scale purification (called “miniprep”). If the E. coli strain is growing fast (such as Mach1) then plasmid can be isolated the same day, otherwise minipreps are incubated overnight and high-quality plasmids are isolated using commercial miniprep kits.

Colonies need to be further screened for the presence of the desired DNA construct and to confirm correct sequence. Several approaches can be used to select potential clone with correct insert. First, to confirm if cloning has been successful, vector can be analyzed using restriction enzymes (several DNA fragments of predicted size should be observed) or by PCR using primers flanking insert site (correct length DNA fragment as analyzed in the gel). To be sure that the inserted fragment is correctly assembled (correct direction) or without additional mutations, fragment sequencing is usually performed. Once confirmed, desired colonies may be employed in downstream applications such as plasmid isolation, subcloning, transfection, and protein expression.

Tips for successful bacterial transformation

Measuring performance and preventing procedural errors are key for better experimental outcomes. Click the accordions for helpful suggestions on setting up controls, calculating transformation efficiency, and preventing arcing.

How to set up positive and negative controls

Positive and negative controls should be included in the transformation step to evaluate the success of the experimental procedure.

ControlSample componentsAssessment
  • Cells alone
  • No DNA
  • Background colony formation
  • Stability of antibiotic(s) in plate
  • Cells
  • Intact plasmid carrying the desired selectable marker (e.g., antibiotic resistance)
  • Cell viability
  • Transformation efficiency
  • Protocol success
  • Cells
  • Recombinant DNA of interest

How to prevent arcing during electroporation

  • Minimize the ionic strength of DNA solutions and electroporation buffers. Ligation DNA mixtures should be column-purified and resuspended in water or TE buffer to remove proteins and salts prior to electroporation.
  • Keep the volume of the DNA solution at no more than 5% of the total cell suspension volume (e.g., 2 µL DNA per 40 µL of cells).
  • Avoid carryover of agar during preparation of electrocompetent cells.
  • Make sure no air bubbles are present in the electroporation cuvette.
  • Dispense the cells directly to the bottom of the cuvette.

Best practices for bacterial transformation

In all steps of bacterial transformation, use sterile tools and labware, media, and reagents as required. It is recommended that once the cells are harvested for further processing, all samples, reagents, and equipment be kept at 0–4°C to improve cell viability and maintain transformation efficiency. Click the accordions for more best practices.

Tube selection for chemical transformation

Polystyrene tubes should be avoided, as DNA can adhere to the surface, reducing transformation efficiency. Traditionally, 17 x 100 mm round-bottom tubes have been used for best results. Using 1.5 mL microcentrifuge tubes may result in poor heat distribution due to smaller surface-to-volume ratios of cell suspension, which can reduce transformation efficiency by as much as 60–90%, especially for the higher-efficiency cells.

Preserve unused competent cells

For storage, it's recommended to aliquot the prepared cells into single-use volumes and store them in screw-cap microcentrifuge tubes. Each freeze/thaw cycle reduces the transformation efficiency by about half. Competent cells should remain stable for approximately 6–12 months when stored at –70°C with minimal temperature fluctuations. Cells should not be frozen or stored in liquid nitrogen, as this practice drastically reduces viability.

Cell plating technique to obtain distinct colonies

Even distribution of the cells on the agar plate is critical for analysis of the colonies. A sterile hockey-stick or L-shaped cell spreader is commonly used to spread the cell suspension while gently rotating the plate (Figures 6, 7A). To achieve better results, it is recommended to prewarm plates and make sure they are free of condensation. Avoid puncturing the agar surface while spreading the cells. Alternatively, autoclaved glass beads (4 mm diameter) may be used to spread the cells. In this approach, 10 to 20 beads are placed on the plate after applying the cell suspension, and the plate is gently swirled so that the cell suspension is spread by the beads (Figure 7B). Cells must be spread quickly before the liquid suspension dries. After spreading, allow the plate to dry before incubating overnight at 37°C in an inverted position.

Step-by-step description on modifying a Pasteur pipette to a L-shaped spreader, using a burner

Figure 6. A Pasteur pipette may be turned into a disposable homemade “hockey stick” or L-shaped spreader.

Distributing cells on agar plates (A) Spreading with a L-shaped rod (B) Spreading with glass beads

Figure 7. Two common plating methods. (A) Spreading with a sterile hockey stick spreader. (B) Spreading with sterile 4 mm glass beads and gentle swirling of the plate.


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