FAQs

Plasmid Subcloning and Ligations

See Technical Resources for more information.

1. What parameters may need to be optimized for successful ligation reactions?
2. What controls are recommended to ensure successful subcloning?
3. What controls can be performed to test the competent cells?
4. What controls can be performed to test the ligation reaction?
5. What controls can be performed to test phosphatase treatment?
6. Are there any special considerations when performing T-cloning?
7. What is blue/white screening and how is it performed?
8. What factors govern the proper selection of host bacterial cell strain?
9. What other methods are available to screen recombinant colonies in addition to blue/white screening?

1. What parameters may need to be optimized for successful ligation reactions?
   A number of parameters may need to be optimized to ensure a successful ligation reaction.
   The ratio of the molar concentration of vector to insert is particularly important, and optimum ratios may vary from 8:1 to as high as 1:16 vector:insert, though generally fall in the range of 3:1 to 1:3. The optimum ratio of vector:insert should be determined empirically for each ligation reaction performed, as differences in insert length and sequence can greatly influence the efficiency of ligation to the same vector backbone. Generally, 10-50ng vector DNA is used per ligation reaction, in a minimum volume (i.e. 10µl).
   The ligation incubation time and temperature may also need to be optimized. In general, blunt-ended ligations are performed at 22°C for 4-16 hours, while sticky-end ligations are performed for 3 hours at 22°C or 16 hours at 4°C. Ligation reactions utilize almost exclusively T4 DNA ligase, but E. coli DNA ligase may be utilized for sticky-end ligations, as the efficiency of blunt-end ligations with this enzyme is quite low. Each ligation reaction generally contains 1-10 Weiss units of high quality ligase. See the T4 DNA Ligase Product Insert (#9PIM180) for additional information.
   The bacterial strain chosen for transformation of the ligation reaction and subsequent propagation of the recombinant plasmid may need to be optimized, as well as the temperature at which the bacteria are grown. For example, ligation of large inserts (>7-8 kbp) into pGEM^® Vectors is facilitated by culturing the transformed bacteria at 30°C instead of 37°C.
   For simple subcloning ligation reactions, subcloning efficiency competent cells may be sufficient for successful cloning, but higher competency cells may be required for more difficult cloning experiments. To maximize the chances of obtaining the clone of interest, the highest competency bacterial cells available should be utilized (>1x10⁸ colony forming units/µg DNA).
   The concentration of antibiotic present for selection of the recombinants may need to be optimized. The following table contains the most frequently used antibiotics and the range of concentrations generally utilized:

   Antibiotic	Working Concentration	Stock Concentration
   Ampicillin (Amp)	50-100µg/ml	50mg/ml in water
   Chloramphenicol (Cm)	20-170µg/ml	34mg/ml in ethanol
   Kanamycin (Kan)	30µg/ml	50mg/ml in water
   Streptomycin (Sm)	30µg/ml	50mg/ml in water
   Tetracycline (Tet)	10µg/ml liquid culture; 12.5µg/ml plates	12.5mg/ml in ethanol

   To decrease background and enhance insert ligation, the vector is dephosphorylated in ligation reactions in which a single restriction endonuclease is utilized to linearize the vector. This inhibits self-ligation and favors insert ligation.
   For successful ligation reactions, both the vector and insert DNA should be very clean. Contaminants can interfere with the ligase and inhibit the reaction if present. The Wizard^® PCR Prep (Cat.# A7170) and DNA Clean-Up (Cat.# A7280) Systems may be utilized to purify both the vector and insert DNA prior to the ligation reaction.

2. What controls are recommended to ensure successful subcloning?
   In order to be certain that a subcloning experiment has been successful, and to troubleshoot if the desired clone is not obtained, appropriate controls should be included with each experiment. Among the controls recommended are controls for:

   • Competent cell efficiency
   • Antibiotic efficacy
   • Ligation efficiency
   • Unligated plasmid background
   • Phosphatase treatment
     These controls are further explained in the following sections.

3. What controls can be performed to test the competent cells?
   In order to ensure that the host cells are competent and not contaminated, three tests are recommended. An aliquot of competent cells should be mock-transformed (i.e., no DNA added but otherwise processed in parallel with the experimental samples). Equal volumes of these cells should be plated on medium with and without antibiotic selection. The selective plate should contain no colonies unless there is a contaminating plasmid or if the antibiotic is not efficacious. Colonies from this plate can be analyzed for the presence of plasmid (e.g., by minipreps or amplification) in order to differentiate these possibilities. The plate without selective medium will have many colonies, or even a lawn of cells, unless the cells are not viable.

   The third test which needs to be performed to test the competent cells is to determine the transformation efficiency of the cells. In order to do this, a known amount of intact plasmid should be transformed into the cells and a known amount of the transformation plated on selective medium. The transformation efficiency of the competent cells in colony forming units (cfu)/µg DNA can then be calculated as follows: After 100µl competent cells are transformed with 0.1ng intact plasmid DNA, the transformation reaction is added to 900µl of medium (0.1ng DNA/1ml), and the bacteria are allowed to recover for a short period of time (1 hour). From that volume, a 1:10 dilution with medium (0.01ng DNA/ml) is made and 100µl plated (0.001ng DNA/100µl). If 200 colonies are obtained, the transformation efficiency is 2 X 10⁸ cfu/µg:

   200cfu/0.001ng=2 x 10⁵ cfu/ng=2 x 10⁸ cfu/µg DNA

   A transformation efficiency of 1 x 10⁶ cfu/µg is sufficient for routine subcloning experiments, 1X 10⁷ cfu/ug is needed for more complex subcloning or transformations with limiting quantities of DNA, and specialized applications such as T-cloning, library construction, or mutagenesis require transformation efficiencies of > 1 x 10⁸ cfu/µg.

4. What controls can be performed to test the ligation reaction?
   In order to test the ligation reaction, linearized (single restriction enzyme-digested, not phosphatase treated) plasmid should be self-ligated in parallel with the experimental samples. An aliquot of this reaction should be transformed and plated on selective medium. The number of colonies on this plate is then compared to an equal amount of cells which have been transformed with an equal amount of linearized, unligated vector and plated on selective medium. The number of colonies on the plate with self-ligated plasmid will be much greater than that of the cells transformed with linear, unligated plasmid if the ligation is sucessful. Moreover, the number of colonies on the linear, unligated plasmid (unligated plasmid background) will be low if the plasmid's linearization was efficient. The number of colonies with the self-ligated plasmid should be comparable to the number of colonies obtained with the uncut plasmid transformation control if the ligation reaction was efficient.

   Alternatively, a quick qualitative test of the ligation can be performed by running, on an agarose gel, a known quantity of unligated plasmid alongside a similar quantity from the ligation reaction. The aliquot from the ligation reaction can be seen to run with a visibly decreased mobility.

5. What controls can be performed to test phosphatase treatment?
   To significantly reduce the background self-ligation of linearized plasmid, the vector can be dephosphorylated prior to the ligation reaction. Generally, linearized plasmid DNA is treated with Calf Intestinal Alkaline Phosphatase (Cat.# M1821) for 60 minutes at 37°C at approximately 0.02 units per pmol DNA ends, followed by inactivation of the phosphatase and purification of the dephosphorylated DNA. Phenol/chloroform/isoamyl alcohol extraction and gel-purification are recommended to purify the dephosphorylated linearized plasmid DNA as CIAP can be difficult to remove from the DNA, and will inhibit subsequent ligation reactions. If the vector is treated with phosphatase prior to ligation, a sample of linearized, phosphatase-treated vector should be self-ligated, transformed and plated on selective medium. The amount of DNA transformed should be the same as that used in the ligation control transformations. When compared to the number of colonies from the ligation control plates, this plate will have approximately the same number of colonies as the linear, unligated plasmid and far fewer colonies than the linear, self-ligated plasmid if the phosphatase treatment was successful and inhibited self-ligation. See the Calf Intestinal Alkaline Phosphatase product insert (#9PIM182) for more information.

6. Are there any special considerations when performing T-cloning?
   Many thermostable DNA polymerases, such as Taq DNA Polymerase and Tfl DNA Polymerase, add an additional nucleotide to the 3' termini of PCR products in a template-independent manner. In general, a single adenine nucleotide is added to the 3' end of the PCR product. Linearized plasmid vectors containing a single T overhang at the 3' end of each strand can then be utilized in ligation reactions, to complement the single A overhang of the PCR product. PCR products generated by thermostable DNA polymerases which possess proofreading 3'-5' exonuclease activity, such as Pfu DNA Polymerase, are blunt-ended products. These products can also be utilized in T-cloning by including an additional 10 minute incubation step at 72°C with Taq DNA Polymerase after the PCR cycles are complete (see Promega Notes 62, 15). In general, T-cloning is more efficient than blunt-ended ligations. To ensure successful and efficient production of recombinant colonies, high efficiency competent bacterial cells (>10⁸ cfu/µg DNA) should be used in the transformation reaction. In addition, high quality DNA Ligase should be used, to avoid nuclease contamination which can cleave off the T or A overhangs. For optimal ligation, the T-cloning reactions should be incubated overnight at 4°C. These conditions are optimal for T-cloning and suboptimal for blunt-ended ligations. For more information on pGEM^®-T and pGEM^®-T Easy Vectors consult Technical Manual 042.

7. What is blue/white screening and how is it performed?
   The pGEM^®-Z Vectors possess a multiple cloning region within the alpha-peptide coding region for the enzyme beta-galactosidase (LacZ gene). Insertional inactivation of the alpha-peptide allows recombinant clones to be identified by color screening on indicator plates. Vectors without an insert express a functional beta-galactosidase enzyme when expressed in bacteria which contain an episomal or chromosomal deletion in the lacZM15 gene, which results in inactivation of the endogenous alpha-peptide. The alpha-peptide from the pGEM^®-Z Vectors or other lacZ containing plasmids complements the omega fragment of beta-galactosidase present in the bacteria. The resulting functional beta-galactosidase enzyme (alpha plus omega fragments) converts substrates such as X-Gal to a colored product, resulting in blue colonies. Cloning inserts into the multiple cloning region of the pGEM^®-Z Vectors disrupts the alpha-peptide coding sequences, and thus inactivates the beta-galactosidase enzyme resulting in white colonies. Small inserts which happen to be in frame with the alpha-peptide coding region may produce light blue colonies, as beta-galactosidase activity is only partially inactivated. Recombinant plasmids are transformed into the appropriate strain of bacteria (i.e. JM109, DH5alpha), and subsequently plated on indicator plates containing 0.5 mM IPTG and 40 µg/ml X-gal. Alternately, 50µl of the X-Gal stock and 100µl of the IPTG stock can be directly added to each plate and the liquid dispersed over the entire plate and allowed to absorb into the agar for 30 minutes at 37°C. In general, IPTG is dissolved in sterile water to a stock concentration of 100mM, and X-Gal is dissolved in N,N'-dimethyl formamide (DMF) to a stock concentration of 50mg/ml. Both the IPTG and X-Gal should be aliquoted and stored at -20°C, and are stable for up to 2-4 months at this temperature.

8. What factors govern the proper selection of host bacterial cell strain?
   The selection of the proper bacterial host cell strain is governed by a number of factors. The bacterial strain selected should not carry endogenous resistance to the antibiotic resistance gene present on the plasmid of interest. If blue/white screening is going to be employed, the host cell strain should possess an episomal and/or chromosomal deletion of the alpha-peptide coding region for beta-galactosidase (i.e., LacZdeltaM15). To propogate M13 and other single-stranded DNA bacteriophages, bacteria must possess a sex pili encoded by an F factor. Examples of bacteria capable of blue-white screening and production of single-stranded DNA upon infection of helper phage include JM109 and DH5alpha^®. A bacterial strain that is deficient in recombination mechanisms may be selected if recombination between the insert and vector or chromsomal sequences is problematic. For expression of recombinant or fusion proteins, a strain of bacteria that is defective in protease activity may be beneficial (i.e., lon mutation present in Y1089 and Y1090). For inducible expression of recombinant proteins, a strain that carries an inducible T7 RNA Polymerase gene are often very useful (i.e. DE3), especially for recombinant proteins which are toxic to E. coli. For mutagenesis experiments, bacteria that are deficient in mismatch repair mechanisms are often utilized to allow replication of the mutated daughter strand (i.e., mutS strains).

9. What other methods are available to screen recombinant colonies in addition to blue/white screening?
   In addition to blue/white screening, recombinant colonies may be screened for the presence of the desired insert by restriction digestion of miniprep plasmid DNA or by direct PCR amplification of bacterial colonies. In general, plasmid DNA from a 1-10ml culture can be isolated and subjected to diagnostic restriction digestions. Comparison of the digestion patterns to uncut miniprep plasmid DNA and to other colonies, particularly blue colonies if available, can verify the presence and orientation of the desired insert. The Wizard^® Plus Miniprep Plasmid DNA Purification Systems may be utilized for isolation of plasmid DNA from mini-prep bacterial cultures (Cat.# A1330, A1340, A1460, A1470, A7100, A7500, A7510). These systems come in either spin or vacuum formats, and the purified plasmid DNA is suitable for many downstream applications, including PCR, restriction digestion analysis, and manual or automated sequencing.

   More recently, direct screening of bacterial colonies using PCR has been developed. In general, a single bacterial colony is added to each PCR tube containing the appropriate amount of water for a 50µl amplification reaction using a sterile pipette tip. The tip is agitated in the water to remove the colony, and subsequently stabbed into an agar grid plate containing the appropriate antibiotic selection, to use as a source of positive clones. Alternately, a single colony can be added to 50µl sterile water, boiled for 5-10 minutes, then centrifuged for 2-3 minutes to pellet the cell debris. An aliquot (5-10µl) is then used as the template in the PCR reaction (see Promega Notes 45, 19). The remaining components are added to the PCR reaction and subjected to normal cycling parameters for the particular primers. If insert orientation, as well as presence, needs to be determined, utilization of a forward vector-specific primer and a reverse insert-specific primer, or visa versa, allows such determination. If only the presence of the insert needs to be determined, then two insert-specific primers can be used. An additional 2-5 minute denaturation step at 94°C before the amplification cycles will aid in lysing the bacteria to enhance PCR product amplification success. The resulting PCR products are then checked on an agarose gel for the presence of the predicted band(s).

   Another method to screen colonies is to perform colony hybridizations with a probe that is know to hybridize to the desired insert. In this method bacterial colonies are either grown on a solid support such as a nylon membrane or transferred to a solid support after growth. The bacteria are lysed in situ and the plasmid DNA is bound to the membrane. The membrane is then probed with a nucleic acid probe. Although colony hybridizations require more initial work than screening with minipreps or PCR, many colonies can be screen rapidly in this manner making it attractive when screening for rare cloning events. Details of colony hybridization techniques can be found in most standard laboratory protocol books such as Molecular Cloning: A Laboratory Manual, Sambrook et al (1989) Cold Spring Harbor Press, Cold Spring Harbor, New York.