Bacterial transformation is a genetic process where bacterial cells take up foreign DNA from the extracellular environment.Because of these newly introduced genes, the bacterial cell is phenotypically altered, or, “transformed.”

Since the DNA uptake occurs from the surrounding environment of the cells, transformation is essentially a horizontal rather than a vertical (i.e., from the parents to the progeny) gene transfer mechanism.

horizontal gene transfer vs. vertical gene transfer in bacterial transformation

Figure 1. Schematic representation of the difference between horizontal and vertical gene transfer. While genes are transmitted from parents to offspring in vertical transmission (left panel), genes are taken up from the surrounding environment or cells in case of horizontal transmission (right panel). [*=Genetic element transfer to the recipient from donor cell or surrounding environment]

Scientists first reported in 1928 that transformation occurs naturally in bacteria under certain circumstances. However, in the laboratory setting, bacteria need to be appropriately treated to make them “competent” to pick up extracellular DNA and get transformed. For a deeper understanding about competent cells, including artificial and natural competence, and how cells are made competent, take a look at this detailed article.

Article Contents

How is bacterial transformation used in molecular cloning?

A quick recap of molecular cloning

Why bacterial transformation is required in the molecular cloning of a transgene?

Bacterial transformation procedure overview

Introducing the vector into the competent recipient cells

Stabilizing freshly transformed cells

Phenotypic lag - the time between recovery and expression

Recovering transformed bacterial colonies

Further screening of transformants to recover the desired colonies

Why do we need to screen transformant colonies?

Colony PCR

Diagnostic/ analytical Restriction Endonuclease (RE) digest

Blue-white screening

Colony PCR vs Diagnostic Digest vs Blue-white Screening


How is bacterial transformation used in molecular cloning?

Bacterial transformation is a critical step in molecular cloning. To understand why researchers use bacterial transformation in molecular cloning, let us first take a very quick recap of molecular cloning.

A quick recap of molecular cloning

A major step in molecular cloning is creating the recombinant vector where the target insert is integrated into the vector molecule at a precise location in a specific orientation.

 two key steps in molecular cloning: A) Creation of the recombinant vector, and its B) Replication and gene expression using a host cell’s physiological processes.

Figure 2. Schematic representation of the two key steps in molecular cloning: A) Creation of the recombinant vector, and its B) Replication and gene expression using a host cell’s physiological processes.


Subsequently, the cloned transgene in this recombinant DNA molecule needs to be:

  1. copied in appropriate numbers, and
  2. expressed by transcription and translation processes.

Now that we are reminded of the fundamental purposes of molecular cloning, let us seek to answer:

Why bacterial transformation is required in the molecular cloning of a transgene?

For achieving both an appropriate number of copies of the recombinant plasmid and for the cloned transgene to be expressed, the physiological machinery of host (most commonly bacterial) cells is used.

Therefore, a vector construct is introduced into a recipient bacterial host by transforming it to exploit the cell’s replication machinery.

The same host may be used for expressing the transgene using the cell’s transcription and translation machinery. Alternatively, often the transgene is shuttled to another suitable host (optimized for gene expression of the recombinant vector) for its expression and downstream analysis.

Therefore, bacterial transformation is a key step in molecular cloning.

Bacterial transformation procedure overview

Five basic steps of the procedure are as follows:

  1. Purchasing commercially available bacterial competent cells or preparing them in the lab.
  2. Introducing the vector into bacterial competent recipient cells.
  3. Stabilizing the transformed recipient cells, letting them recover from the stress of the transformation procedure.
  4. Selecting transformed colonies that have the vector.
  5. Ensuring the transformed bacterial colony contains the transgene in the correct sequence orientation.

Details of steps 2 -5 are described below. A schematic representation of bacterial transformation is visualized below.

bacterial transformation protocol outline

Figure 3. Schematic outline of bacterial transformation

Introducing the vector into the competent recipient cells

As shown in Figure 3, recipient host cells can be transformed either via electroporation or using heat-shock method depending on whether they are electrocompetent or chemically competent. Underlying principles of each method are described in this article.

Transformation by electroporation

The electroporation method relies on passing electric current, in the range of 1000 volts, for up to a millisecond duration through the liquid suspension of the competent cells and vector mixed together.

Figure 3 depicts a schematic representation of electroporation. The cuvette holding the competent cell-vector mix is put between the two electrodes of the electroporation apparatus and an electric pulse is applied.

The cell membranes, unable to pass current, act as a capacitor, which results in the phospholipid bilayer temporarily changing architecture giving rise to pores through which the vector molecules can get inside the cell from the extracellular milieu.

how electroporation works

Figure 4. Illustration of how electroporation works. After a current is sent, the phospholipid bilayer is restructured leading to openings in the cell membrane allowing plasmid entrance.

Experimental tip:Ensure that no air bubbles form in the cuvette while pipetting the recipient cell-vector mix into it. Air bubbles cause arcing. If that happens, it is highly unlikely that the procedure will work and viable transformants will be recovered at the next steps. If you have struggled with arcing in the past or want to learn more about it, check out our troubleshooting guide.

Transformation by heat-shock method

Inducing chemical competence followed by heat-shock assisted transformation of recipient cells is a physicochemical procedure.

While treatment with cations, most commonly Ca2+, comprises the chemical aspect, shock fromdrastic temperature fluctuations (from 0 degrees to 42 degrees Celsius) is the physical part.

The exact underlying mechanisms of this method of transformation is not very clearly described in literature. Nevertheless, the current understanding of the putative contributions of both the cations and the temperature shock are described below.

Indeed, the chemical treatment appears to induce “competency” in the recipient cells while the temperature shocks facilitate the actual internalization of the foreign DNA.

How do divalent cations such as Ca2+ help in bacterial transformation?

  • Both the extracellular DNA and the recipient cell membrane are negatively charged, and repel each other. Divalent cations neutralize these negative charges thereby reducing the repulsion (Figure 5).
  • Ca2+ bound DNA adsorb onto the recipient cell surface more easily.
  • Ca2+ , upon binding to the cell membranes, promotes permeability.
  • Divalent cations promote intramolecular static attractive forces in the extracellular DNA. This induces the vector DNA to adapt a compact ball-like geometry which readily shuttles in through the membrane pores of the recipient cells.

Role of CaCl2 in bacterial transformation.

Figure 5. Role of CaCl2 in bacterial transformation. Before treatment, the negative charges between the cell and plasmid leads to repulsion. Treatment with Ca2+ neutralizes the charges.

How does temperature shock aid in bacterial transformation?

  • Prolonged incubation on ice (Figure 3) stiffens the lipid structures in the recipient cell membranes resulting in loss of their fluidity. This stabilizes the Ca2+-cell membrane interactions.
  • The high temperature (420C) heat shock is thought to increase the Brownian motion in the extracellular environment. Just to clarify, Brownian motion is random particle movement within fluid. This helps pushthe external vector molecules to the interior of the cells.
  • The transition from 00C to 420C leads to lipid loss in the cell membrane, while the subsequent drop to 00C again causes protein loss. So, the application of cold-heat-cold shock is critical in this procedure to facilitate creation of permeating pores by depleting the cell membranes of both proteins and lipids. Further, in this process, depolarization of the membrane occurs, reducing the repulsion between similarly charged vector DNA and host cell membranes.

Stabilizing freshly transformed cells

Both methods of transformation cause significant stress on the cells. So, it is necessary to transfer bacterial cells to a growth-promoting environment immediately following transformation.

This is done by suspending the bacteria in 1ml of pre-warmed rich liquid media such as LB right after the last step.

Since common recipient bacteria, usually E. coli, used in the lab setting are mesophilic and aerobic, the culture is shaken at 370C for 30-60 minutes (Figure 6).

Phenotypic lag - the time between recovery and expression

As we discuss this idea of recovery and stabilizing your competent cells, we need to introduce you to the concept of phenotypic lag.

Here’s what happens: after the vector is introduced into the cell, there is a lag or a period of time between introduction of the vector genes in the recipient cell and the final gene products, i.e., the corresponding proteins, being produced.

Therefore, your suspension is going to have an interesting mix of bacterial cells. Some did not get genetically transformed because, for whatever reason they did not take up the vector. The genetically transformed cells have taken up the vector; however, the vector genes (including the selectable marker gene(s) such as those conferring resistance to antibiotic(s)) have still not started expressing the corresponding proteins. Thus, at this stage, ie, right after transformation, these cells are not phenotypically transformed. Little bit of time is required for the genetically transformed cells to start expressing the vector genes and thus get phenotypically transformed. This time difference is known as “phenotypic lag”.

In figure 6, we detail this concept of phenotypic lag and phenotypic transformation. Notice the cell in orange (in the microcentrifuge tube) has been genetically transformed, but is not yet phenotypically transformed. The orange cell in the test tube is both genetically and phenotypically transformed.

phenotypic lag

Figure 6. Schematic illustration of phenotypic lag.

Notwithstanding salvaging and stabilizing the bacterial cells from the transformation stress, this step accounts for the time taken by the newly transformed cells to express the genes encoded by the vector (most commonly plasmid).

This is critical because the next step involves selecting and recovering the transformants out of the entire competent cell population by virtue of the selectable marker gene on the vector. Typically, antibiotic-resistance protein encoding genes are used as selectable markers.

In other words, if the bacterial cells are plated on antibiotic-containing (selection pressure) solid media right after transformation, both transformed (containing the vector, and therefore the antibiotic resistance gene) and non-transformed (no vector, and thus no antibiotic resistance gene) cells would not survive (Figure 6).

Recovering transformed bacterial colonies

As shown in figure 6, after accounting for the phenotypic lag, the bacteria are plated on selective solid media.

Most commonly, selectable markers on plasmids encode resistance to a particular antibiotic. Consequently, as a selection pressure, the media contains the corresponding antibiotic. Only the transformed cells grow (positive selection) as colonies. The non-transformed bacteria die (and are thus selected out) because of the antibiotic pressure.

how antibiotic selection works for bacterial transformation (selection pressure)

Figure 7. A closer look at how selection pressure works. The plasmid being introduced has an antibiotic resistance gene for ampicillin. In the suspension, some cells took up the plasmid (depicted in green), while the others did not (depicted in blue). All of the cells were plated on media containing ampicillin. But only the bacterial cells with the plasmid grew (depicted as green colonies).

Experimental tip: First, it’s best to plate 50µL from the 1ml culture onto the first antibiotic plate; the remaining 950 µL should be spun down using a table-top centrifuge, resuspended in 50-100 µL LB and plated onto a second antibiotic plate. In case the transformation efficiency is high, plating the whole of the 1ml culture onto a single plate might produce a lawn growth without discernible isolated colonies. Conversely, if the transformation efficiency is low, the first plate (50µL of culture plated) would have no colonies while the second plate (950µL of culture plated) would have a few colonies. Second, it’s good practice to streak out even single isolated colonies a second time on a fresh antibiotic plate. Sometimes two or more colonies stick to each other giving a false impression of a big single colony. Obtaining single isolated colonies is a necessity in this procedure.As you will see in the next step, not all transformants have the right construct despite having the vector backbone which confers the antibiotic resistance phenotype. Further, due to antibiotic degradation in localized areas of the plate, small satellite colonies emerge which are antibiotic-sensitive vector-negative bacterial populations. Streaking out a second time gets rid of these as well.

We have a transformation efficiency calculator that you can use.

Further screening of transformants to recover the desired colonies

While the previous step selects in the transformed cells by eliminating the non-transformed cells by using selection pressure (antibiotic), further screening of the transformants is indeed required to identify the bacterial colonies that have the desired recombinant vector construct. Details are described below.

Why do we need to screen transformant colonies?

If the ligation mix is directly used as a source of the recombinant vector DNA in the transformation step, it would have all these DNA molecules in varying proportions:

  1. insert fragment
  2. linearized vector
  3. self-circularized (by autoligation) vector
  4. correct recombinant vector with the insert ligated within it at the correct position and sequence.

outcomes of bacterial transformation - bateria will take up linearized plasmid with insert, no plasmid, insert only, full plasmid

Figure 8. Shows all the possible outcomes of transformation. Here, cells could have taken up other components besides the correct plasmid. These components include only the fragment, the linearized plasmid, a recircularized plasmid without the insert, and the correct recombinant plasmid.

Further, if a multiple insert cloning reaction is involved, for example, using Gibson assembly method, then there might be recombinant vectors with some but not all fragments cloned in proper sequence.

Now let’s analyze the outcome of transformation using each of these DNA molecules.This is depicted in figure 9.

  • Recipient cells transformed with insert fragment (1) and/or linearized vector (2) would not survive the selection pressure when plated on solid media because a circular vector is required to express the selectable marker.
  • Cells with blank (no insert) self-recircularized vector (3) would produce colonies on the antibiotic plate.
  • Cells with recombinant vector having the cloned transgene (4) would also produce colonies on the antibiotic plate. Transformants with possible variants of the desired recombinant vector (consider a multi-fragment cloning experiment where only some but not all insert fragments have been cloned in the vector) will also produce colonies.

bacterial transformatin plated with different plasmid scenarios. Shows which transformed colonies will grow and which ones did not.

Figure 9. Shows what happens on a plate with antibiotic. The cells with just the insert and the cells with the linearized plasmid will not survive antibiotic selection pressure. Cells with the recircularized vector (containing genes for antibiotic resistance) and cells with the correct plasmid will survive the selection pressure on the plate.

Please note that transformants with blank (no insert) self-circularized vector or the correct recombinant would produce colonies on the antibiotic plate. Just to clarify, the antibiotic resistance gene is found on the vector itself, and not the insert. This means cells with blank recircularized plasmid (scenario 3 above in figure 9) and those with recombinant plasmid (scenario 4 in figure 9) would survive on the antibiotic selection plate.

Therefore, to discern between all these different transformant variants and to screen only the correct colonies, at least one of these subsequent diagnostic steps is necessary.

Colony PCR

In colony PCR, a single isolated transformant colony is directly used rather than the plasmid being purified out of the bacteria as the source of template DNA in a PCR reaction.

When the PCR thermocycler heats up the cells in the PCR mix, it helps lyse the cells, releasing the vector DNA into the reaction. A schematic representation of this method is depicted in Figure 10.

 colony PCR. Colonies are aseptically picked using a pipette tip

Figure 10. Schematic representation of colony PCR. Colonies are aseptically picked using a pipette tip or toothpick, patched onto a replica plate and are put into the PCR reaction as the source of template DNA. The bands produced by different colonies are analyzed on a DNA gel. Here, the PCR reveals that colonies 1 and 3 contain the empty plasmid as they produce the same band as the control (C) (empty vector). Colony 4 contains the correct recombinant plasmid construct with the insert. In Colonies 2 and 5 PCR did not work for some reason, producing only a primer dimer.

Experimental tip: The colony is picked up from the agar plate using a sterile toothpick or pipette tip and is first patched onto a second plate before being added into the corresponding PCR tube (Figure 10). This serves the following two purposes: Too much bacterial cells in the PCR mix often is detrimental to the reaction, producing smudged unclear bands. Patching onto a second plate reduces the cell load going into the PCR tube. For ensuring aseptic microbiology practice, a colony where a pipette tip or toothpick (albeit autoclaved) has touched should not be again used to start a culture (in case it indeed is confirmed by PCR to be a desired strain) and store it. The second plate which now has the replica of this colony can be used for this purpose.

When choosing the primer pair, one or many of the following 3 options can be chosen (Figure 11):

  1. Primers that bind at the two ends of the insert fragment (Insert-specific). The colony that has the insert integrated within the plasmid produces a band on the DNA agarose gel, while the ones with only the blank vector produces no band. However, in this case it would be difficult to differentiate between the cases of a true negative colony (blank plasmid) versus the PCR not working for a colony for any reason.
  2. The above problem can be overcome if primers bind to the plasmid backbone just upstream and downstream of the insert cloning site (backbone-specific). The colonies with empty vectors produce shorter bands compared to those with the insert.
  3. Primers may be designed to ensure that the insert has been integrated within the vector in a specific orientation (orientation-specific). This is especially helpful in cases where the insert-vector cloning lacks directionality (for example, blunt-end cloning or when only one RE is used to digest the to molecular cloning article).

how to choose a primer for colony pcr and things that can go wrong

Figure 11. Choosing primers for a colony PCR. If insert specific primers are used (left most panel), then for the colony with blank plasmid the primers fail to bind at all (red cross sign) and thus no band is produced (only fuzzy very low molecular weight primer dimer), while the correct colony (having the desired recombinant plasmid) will produce a band corresponding to the size of the insert. For backbone specific primers (central panel), the correct colony (having the desired recombinant plasmid) will produce a higher molecular weight band (comprising of the insert size plus upstream and downstream sequences where the forward and reverse primers bind on the plasmid backbone) than the colony with the blank plasmid; difference in band size for these two colonies is the size of the desired insert fragment. The right most panel depicts the use and gel electrophoresis pattern when orientation-specific primers are used. The correct colony (having the desired recombinant plasmid where the insert has been cloned in the correct orientation) will produce a band; the colonies with either blank plasmid (primers fail to bind at all, as depicted by red cross sign)or where the insert has been cloned in the flipped orientation will produce only primer dimers on the gel. [“+” sign denotes colony with desired construct while “-” sign denotes undesired colonies either lacking plasmid or having the wrong construct].

Diagnostic/ analytical Restriction Endonuclease (RE) digest

Few of the transformant colonies are grown up as cultures, and the corresponding vectors are isolated from them (Figure 12). In the next step, the vectors are digested by one or more restriction endonucleases The fragments thus formed are analyzed using electrophoresis. As shown in figure 12a, the plasmid constructs harboring the insert produces a different band pattern on the gel as compared to the blank plasmid.

diagnostic plasmid digest to confirm presence of the cloned transgene

Figure 12. Schematic representation of diagnostic plasmid digest to a) confirm presence of the cloned transgene, and b) confirm orientation of the insert fragment.

a) 900 bp in the MCS of the empty plasmid (5000bp in size) is replaced with 1200 bp of the insert, thereby producing the recombinant construct. If the blank plasmid is digested with Restriction enzyme (RE) 1, it gets linearized producing a single band of 5kb (lane 2 of the gel from the left). If it is digested with RE1 and RE2, two fragments (corresponding to the two bands on lane 3 of the gel) are produced: 4100bp and 900bp. If the recombinant plasmid (containing the 1200bp insert) is digested with RE1 and RE2 and run on the same gel, two bands measuring 4100bp and 1200bp are produced (last lane in the gel). This allows differentiation between the blank plasmid and the recombinant plasmid construct.

b) If the cloning process lacks directionality (for example because of using single Restriction enzyme digestion or blunt-end cloning etc.), the insert may be cloned in the correct or incorrect (exactly flip of the correct) orientation. To distinguish between these two possibilities the corresponding plasmid constructs are digested with the same restriction enzymes (RE1 and RE2 in this case). Because of the specific sites where the enzymes cleave, the corresponding bands on the gel are different, thereby allowing differentiation and selection of the correctly cloned insert.

Figure 12b depicts how this method can be used to monitor directionality of cloning. If the insert is in one orientation, a characteristic RE-digest pattern is seen on the gel which is quite different if the orientation is reversed.

Confirming the directionality of cloning of the transgene is very important. If the transgene is not cloned in the proper orientation, it will not express the downstream gene product.

Blue-white screening

Blue-white screening is a method where positive colonies containing the insert in the plasmid construct can be readily identified visually and contrasted from the colonies transformed with empty vectors.

The schematic outline of this screening method is depicted in Figure 13. An underlying enzymatic mechanism exploited herein is as follows. The lacZ gene encodes the enzyme betagalactosidase which can cleave X-gal into galactose and a blue colored compound 4-chloro 3-bromo indigo.

As evident from figure 13, a specialized plasmid-bacterial strain combination is exploited in this screening. The recipient strain is deficient in producing the LacZ enzyme. The corresponding plasmid vector encodes this protein within its multiple cloning site (MCS).

Thus, transformants, which have the blank plasmid and hence the full lacZ gene within the uncut MCS produce blue colonies on the antibiotic plate containing X-gal.

If the insert is cloned in the MCS of this plasmid, the lacZ gene is cleaved and no longer expresses the betagalactosidase enzyme. Consequently, colonies that have the insert cloned within the vector are white.

plasmid has the lacZ gene within its MCS. Both this plasmid and the insert are digested with the same pair of restriction enzymes

Figure 13 a) The specialized plasmid has the lacZ gene within its MCS. Both this plasmid and the insert are digested with the same pair of restriction enzymes (RE1 and RE2), thus producing corresponding complementary overhangs (depicted by sky-blue and violet colors)

b) The digested plasmid fragments and insert are there in the ligation reaction. Here two outcomes are possible: i) the plasmid may self-circularize using the digested (cut out) plasmid fragment containing the lacZ gene which has sticky overhangs complementary to the plasmid backbone ii) the insert may ligate with the plasmid backbone owing to its complimentary sticky ends, thus creating the recombinant vector

c) Competent bacteria are transformed directly with the ligation mixture which has both plasmids i and ii. Hence there will be two kinds of transformants possible: one which contain plasmid i, and the other which contain plasmid ii.

d) Since both these plasmids contain the functional antibiotic resistance gene (AbR), both types of transformants will survive on the antibiotic plate. However, only the one with the self-circularized plasmid (i) will produce blue colonies due to expression of the betagalactosidase enzyme (which cleaves X-gal to produce blue color metabolite) due to the presence of the functional lacZ gene; the second type of transformant colonies will produce regular white colonies because the recombinant plasmid (ii) they harbor does not have the lacZ gene

Colony PCR vs Diagnostic Digest vs Blue-white Screening

  • Time consumption: The blue-white screening method allows ready visual differentiation between negative and positive colonies. The diagnostic digest takes the maximum time because the additional steps of growing colonies as liquid cultures and isolating plasmids from them are involved.
  • Cost: The diagnostic digest is most costly, due to the involvement of multiple restriction endonuclease enzymes. IPTG, the chemical used to switch on the lac operon, and X-gal are not comparably expensive. For the PCR method, a relatively lost cost Taq DNA polymerase can be used; since the PCR fragment is not being used for downstream steps (such as cloning), a costly high-fidelity DNA polymerase is not required.
  • Effectiveness: All three methods can confirm the presence/ absence of the insert in the plasmid construct harbored by the transformant colonies. However, blue-white screening is unable to confirm the orientation of the insert, which the other two methods can, if the experiment is properly designed (Figures 11 and 12). Further, in case multiple insert fragments need to be cloned in tandem (consider Gibson assembly), the blue-white method cannot confirm whether all or only a subset of the required fragments has been cloned in a colony.

In spite of best practices taken during the cloning procedure, mutations may occur in the cloned insert fragment.

Often these are as small as single-nucleotide mismatches; nevertheless, such sequence errors might alter the reading frame or sequence of the downstream gene product. To account for such scenarios, the final step is to confirm the sequence of the final cloned fragment of interest by commercial DNA sequencing.

bacterial transformation workflow

Figure 14. Bacterial transformation process workflow


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Note to self - format in commercio to bring attention.