Molecular Cloning Guide

Agarose Gel Electrophoresis of DNA

Running double-stranded, linear DNA (like plasmid DNA from restriction enzyme digests) on an agarose gel is a routine activity in molecular biology laboratories. The basic method is very straightforward:

1. Set up the gel electrophoresis apparatus as recommended by the manufacturer.
2. Weigh the required amount of agarose and add it to the appropriate volume of TAE or TBE 1X Buffer in a flask or bottle. For example, to prepare a 1% agarose gel, add 1.0g of agarose to 100ml of buffer. Note: The volume of buffer and agarose should not exceed half the volume of the container.
3. Heat the mixture in a microwave oven or on a hot plate for the minimum time required to allow all the agarose to dissolve. Interrupt the heating at regular intervals and swirl the container to mix the contents. Do not allow the solution to boil over.
   CAUTION: The container and contents will be hot! Swirling may also cause solution to boil vigorously. Use adequate precautions.
4. Cool the solution to 50–60°C and pour the gel. Allow the gel to form completely (typically, 30 minutes at room temperature is sufficient). Remove the comb from the gel, place it in the electrophoresis chamber and add a sufficient volume of TAE or TBE 1X buffer to just cover the surface of the gel.
5. Load samples with 1X Blue/Orange Loading Dye into the wells.
6. Connect the gel apparatus to an electrical power supply and apply an appropriate voltage to the gel. Standard gel electrophoresis of DNA fragments greater than 2kb typically call for between 5 - 8 volts/cm. Higher voltages and shorter runs will decrease the resolution of the gel and may also cause overheating that can melt the agarose. 
7. After electrophoresis is complete, remove the gel for staining. Either soak it in a solution of 0.5µg/ml ethidium bromide for 30 minutes at room temperature, or in a solution of Diamond Nucleic Acid Dye diluted to 1:10,000 in the same type of buffer used to cast the gel for 15 to 30 minutes, protected from light. Note: Ethidium bromide or Diamond Nucleic Acid Dye may also be incorporated in the gel during preparation, and in the electrophoresis buffer, at the same concentration as would be used for staining. This eliminates the need for post-electrophoretic staining but may interfere with accurate size determination of DNA fragments. CAUTION: Always wear gloves when working with ethidium bromide.
8. Place the gel on a UV lightbox (if using either stain), or on any other gel documentation system, or blue light transilluminator if using Diamond Nucleic Acid Dye. Photograph the gel according to the specification recommended for your camera. Diamond Nucleic Acid Dye may be visualized and photographed with any system that has a SYBR® Gold filter or setting.  CAUTION: Use protective eyewear when working with a UV light source.
   Note: If using ethidium bromide, you may wish to destain or rinse the gel in fresh 1X running buffer prior to viewing it on the UV lightbox.  
   

Solution Compositions

Nearly all of these reagents can be purchased premade including the agarose gels. Here are the directions if you wish to prepare your own reagents.

Blue/Orange Loading Dye, 6X (Cat.# G1881)
10mM Tris-HCl, pH 7.5
50mM EDTA
15% Ficoll® 400
0.03% bromophenol blue
0.03% xylene cyanol FF
0.4% orange GOne or more dyes can be left out of the recipe to create a custom loading dye.

TAE 50X Buffer (1L) (Available in a 10X or 40X solution from Promega [Cat.# V4271 and V4281, respectively]): Dissolve 242g Tris base and 37.2g disodium EDTA dihydrate in 900ml of deionized water. Add 57.1ml glacial acetic acid and adjust the final volume with water to 1 liter. Store at room temperature or 4°C.

TBE 10X Buffer (1L) (Available in a 10X solution from Promega [Cat.# V4251]): Dissolve 108g of Tris base and 55g boric acid in 900ml deionized water. Add 40ml 0.5M EDTA (pH 8.0) and increase the final volume to 1L. Store at room temperature or 4°C.

Polymerase chain reaction cloning (PCR cloning) allows you to amplify a DNA fragment of interest without the use of restriction enzymes. When PCR was in its infancy, researchers found that cloning PCR products by simple blunt-ended ligation into blunt-ended plasmid cloning vectors was not easy. Part of the challenge is thermostable DNA polymerases, like Taq DNA polymerase, add a single nucleotide base extension to the 3´ end of blunt DNA in a template-independent fashion (Clark, 1988. Mole et al, 1989). These polymerases usually add an adenine, leaving an “A overhang”.

In the evolution of molecular cloning techniques, researchers have developed several methods to address the challenges posed by the presence of A overhangs on PCR products. One method involves treating the product with the Klenow fragment of E. coli DNA Polymerase I to convert these overhangs into blunt-ended fragments suitable for cloning. However, this technique is not particularly efficient for PCR fragments. Another method frequently used by researchers is to add restriction enzyme recognition sites to the ends of the PCR primers to engineer specific cleavage sites within the PCR product (Scharf et al, 1986). This strategy facilitates the straightforward digestion of the PCR product, allowing for its subsequent integration into a plasmid vector. Care must be exercised in primer design when using this method, as not all REs cleave efficiently at the ends of DNA, and you may not be able to use every RE you desire (Kaufman and Evans, 1990). Some REs require extra bases outside the recognition site, adding further expense to the PCR primers as well as risk of priming to unrelated sequences in the genome.

Recent advancements have shifted focused towards more streamlined and efficient cloning methods. The most popular method for cloning PCR products is T-vector cloning (or TA cloning). In essence, the plasmid cloning vector is treated to contain a 3´T overhang to match the 3´A overhang of the amplicon (Mezei and Storts, 1989). The A-tailed amplicon is directly ligated to the T-tailed plasmid vector with no need for further enzymatic treatment of the amplicon other than the action of T4 DNA ligase. We provide T-vector systems based on this technology for routine cloning and direct mammalian expression.

*All bases may be found at 3´ overhang; adenine tends to be encountered most often.

T-Vector Systems:  pGEM®-T and pGEM®-T Easy Vectors

The pGEM®-T and pGEM®-T Easy Vector Systems are designed as simple cloning vectors. The vectors provide convenient T7 and SP6 promoters that can serve as binding sites for sequencing primers, or for promoting in vitro transcription on either strand of the insert when the appropriate RNA polymerase is used. The vectors have the lacZα gene, allowing easy blue/white screening for recombinants using an appropriate bacterial strain (e.g., JM109, DH5α™, XL1 Blue, etc). The pGEM®-T Vectors are provided with 2X Rapid Ligation Buffer,  which allows efficient ligation in just 1 hour with the supplied T4 DNA Ligase. You can either supply your own favorite E. coli cells or purchase the system with Promega JM109 Competent Cells. For mammalian cells, the pTARGET™ Vector is ideal for both cloning and expression of PCR products.

PCR Cloning Controls

Each Promega PCR cloning system is provided with a control insert. The ligation and subsequent transformation of this positive control can give you a lot of information with regard to the ligation and transformation of your insert. The total number of blue colonies obtained with the Promega-supplied positive control insert and no-insert controls should be approximately equal. The negative control may have some white colonies as well.

Interpreting Results from T-Cloning Experiments

Experimental insert looks like control insert in efficiency and percent white colonies: Successful experiment. Greater than 80% of the white colonies should contain inserts.

Experimental insert and control insert look like negative control: Ligation has failed. Avoid multiple freeze/thaws of the ligation buffer. Ligase buffer contains ATP and could be damaged by freeze/thaw cycles. You may need to dispense the ligase buffer into smaller aliquots for your experimental needs.

No colonies with experimental insert, control insert or negative control: Transformation has failed. Reassess the competent cells with an intact, supercoiled plasmid and determine the transformation efficiency. Use cells >1 × 10⁸ cfu/µg to ensure >100 colonies from the control insert ligation.

Experimental insert has more blue colonies than control insert or negative control and fewer white colonies than control insert: In-frame insertion, no interruption of α-fragment.
The pGEM®-T Vector Control DNA will produce recombinants that generate white colonies because the lacZ reading frame is interrupted. White colonies also form with successful insertion of other DNA fragments that disrupt the lacZ reading frame. Occasionally, a successfully inserted fragment does not disrupt the lacZ reading frame, leading to the growth of blue colonies. Although this tends to occur most frequently with PCR products of 500bp or less, inserts of up to 2kb may result in blue colonies. Moreover, some insert DNAs can also result in pale blue colonies or “bullseye” colonies with a blue center and a white perimeter. These are still successful insertions.

Properties of E. coli Cloning Strains

Common laboratory strains of E. coli, like JM109, DH5α™ and XL-1 Blue, are different from their wildtype counterparts. These strains carry some mutations designed to help propagate plasmids. Typically, laboratory strains have a mutation in the recA gene (recA1), a gene involved in recombination. The mutant gene limits recombination of the plasmid with the E. coli genome so that the plasmid inserts are more stable (the recA1 mutation is more effective than the recA13 mutation). Each of these strains also carries the endA1 mutation that inactivates a nuclease that might co-purify with plasmids during purification. This mutation allows for purification of higher quality plasmids. Special treatments must be performed on plasmids from strains that do not have this mutation (e.g., RR1, HB101, etc.) to eliminate the nuclease from the plasmid prep (e.g., an Alkaline Protease digestion).

Common laboratory strains of E. coli are typically defined as K strains or B strains based on the presence of the restriction and modification system that functions around EcoKI or EcoBI, respectively. In a wildtype K strain, the E. coli will have both the EcoKI restriction enzyme to cleave foreign DNA and EcoKI methylase to protect and mask host DNA recognition sequences. In B strains, the EcoBI restriction enzyme and methylase serve the same purpose. Strains like JM109, DH5α™ and XL-1 Blue are K strains but carry the hsdR17 (rK–, mK+) mutation. This mutation knocks out the EcoKI restriction enzyme but leaves the methylase intact. Therefore, these strains will not degrade plasmid DNA isolated from a B or K strain but will methylate it. This is useful if the DNA must be transferred to a K strain with an intact K restriction and methylation system.

If you wish to incorporate blue/white selection into your cloning scheme, you need to transform E. coli carrying a lacZ∆. This mutation deletes a portion of the β-galactosidase gene, leaving what is termed the ω-fragment. The plasmid vector supplies this deleted portion, or α-fragment. Once inside the bacterium, the plasmid produces the α-fragment and the E. coli produces the ω-fragment, which combine to make a functional β-galactosidase. If grown on plate containing 5-bromo-4-chloro-3-indoyl-β-D-galactopyranoside (X-gal), the colony will turn blue as a result of β-galactosidase activity and indicate full complementation of the bacterium by the plasmid. This is termed α-complementation. Blue/White cloning methods use plasmids with a multiple cloning region within the coding sequence of the α-fragment. Disruption of the reading frame due to the presence of the insert will produce a non-functional α-fragment incapable of α-complementation. These disrupted plasmids are differentiated from the plasmids without insert by the color of the colony (white versus blue), hence the term blue/white selection. Strains like JM109, DH5α™ and XL-1 Blue have the necessary deletion. One difference between these strains lies in how you get the bacterium to produce the ω-fragment. Both JM109 and XL-1 Blue have a second mutation call lacIq. This mutation leads to increased production of the LacI repressor that stops transcription from the lac operon until substrate is present. To relieve this repression, these strains are grown on media containing the non-cleavable lactose analog, isopropyl-β-D-thiogalactopyranoside (IPTG). DH5α™ does not have the lacIq mutation and constantly produces a low level of the ω-fragment through leaky transcription of the lac operon and therefore does not require IPTG for blue/white selection.

Making your own competent cells

Preparation of Competent Cells: Modified RbCl Method

This rubidium chloride protocol gives better transformation efficiencies than the CaCl₂ procedure, for most strains. The procedure is an adaptation of one described in Hanahan, D. (1985) In: DNA Cloning, Volume 1, D. Glover, ed., IRL Press, Ltd., London, 109.

Materials to Be Supplied by the User

• LB medium and plates
• LB + 20mM MgSO₄
• TFB1, ice-cold
• TFB2, ice-cold
• dry ice/isopropanol bath or liquid nitrogen

1. Inoculate 2.5ml of LB medium in a plating tube with a single colony from an LB plate. For JM109, use M9 + B1 plate so that the F´ episome is maintained. Incubate the tube overnight at 37°C with shaking (approximately 225rpm).
2. Subculture the overnight culture at a 1:100 dilution. Inoculate 250ml of LB supplemented with 20mM MgSO4 in a 1L flask, with 2.5ml of the overnight culture. Grow the cells at 37°C with shaking until the OD600 reaches 0.4–0.6 (usually 5–6 hours, but the time may vary).
3. Pellet the cells by centrifugation at 4,500 × g for 5 minutes at 4°C. For a 250ml culture, use two 250ml centrifuge bottles in a large rotor; half of the culture in one bottle, half in the other. Be sure to balance the bottles.
4. Gently resuspend the cell pellet in 0.4 of the original volume of ice-cold TFB1. For a 250ml subculture, use 100ml total of TFB1 (50ml/bottle centrifuged). Combine the resuspended cells in one bottle. For the remaining steps, keep the cells on ice and chill all pipettes, tubes and flasks.
5. Incubate the resuspended cells on ice for 5 minutes at 4°C.
6. Pellet the cells by centrifugation at 4,500 × g for 5 minutes at 4°C.
7. Gently resuspend the cells in 1/25 original volume of ice-cold TFB2. For a 250ml subculture, use 10ml of TFB2.
8. Incubate the cells on ice for 15–60 minutes, and then dispense 100µl/tube for storage at –70°C. During dispensing, keep the cells as cold as possible. Flash-freeze the tubes in a dry ice/isopropanol bath or liquid nitrogen, and store at -70°C. JM109 competent cells prepared by this method are stable for 1 year if never thawed.

Note: Competent cells may be conveniently quick-frozen using ice bath racks, or ChillBlock™. Set up an ice bath and an ethanol bath. Label the tubes on their lids. Open the labeled tubes and place them in a rack in the ice bath. Dispense 100µl cells per tube, then close the tubes. Add dry ice to the ethanol bath, wait for it to stop bubbling, and then transfer the rack and tubes to the dry ice bath for about 15 seconds. Regular ice may be used in the ethanol bath if dry ice is not available, but flash freezing will not occur. Transfer the tubes to their storage box and place in a –70°C freezer. Do not get alcohol on the lips of the tubes. Liquid nitrogen also can be used for flash-freezing, but be sure it is compatible with the rack you are using. Use only plasticware designed for liquid nitrogen.

Note: Be careful not to get alcohol on the labels because it will remove them.

Many E. coli strains carry episomes (e.g., F´ and P2) expanding the capabilities of the bacterium for use in cloning applications. For example, the XL1-Blue and JM109 strains carry the lacIq∆M15 mutation on the F´ episome. The episomes are extrachromosomal replicating plasmids with a selectable marker. When making competent cells from strains with episomes, the bacteria must first be plated on selective media. For XL1-Blue, colonies are selected on tetracycline plates because the episome contains the tetR gene. This means that the strain cannot be used with cloning plasmids containing the tetR gene for selection. JM109 cells should be selected first on M9 minimal media containing thiamine (vitamin B1). The bacterial chromosome lacks the biosynthetic genes for proline synthesis (proAB) but the episome carries those genes. Colonies grown on the M9 + B1 plates (recipe on page 48) can then be processed into competent cells ready for blue/white selection. 

Determining Transformation Efficiency of Competent Cells

This is a general protocol for use with the procedure for producing competent cells provided above. Please follow manufacturers’ instructions when using purchased competent cells. Keep in mind, transforming more than 10ng of DNA from a ligation reaction may actually decrease transformation efficiency.

1. Thaw a 100µl aliquot of competent cells on ice.
2. Transfer 100µl of the cells to a 17 × 100mm polypropylene tube prechilled on ice.
3. Add 0.1ng of a supercoiled plasmid [e.g., pGEM®-3Zf(+) Vector] in a 10µl volume to the competent cells and gently mix by swirling the pipet tip (do not mix by pipetting).
4. Transfer the tubes from ice to a 42°C water bath and heat shock for 45–60 seconds. Place on ice immediately to cool for at least 2 minutes.
5. Add 890µl of SOC medium (giving a concentration of 0.1ng DNA/ml) and incubate for 45 minutes at 37°C with shaking (~150rpm).
6. Dilute cells by transferring 100µl of cells to 900µl of SOC medium (giving a concentration of 0.01ng DNA/ml). Plate 100µl of diluted cells onto LB plates with the appropriate antibiotic. You may wish to plate 100µl of undiluted cells for determining efficiency as well. 
7. Incubate the plates overnight in a 37°C incubator and count the number of colonies obtained.

Transforming Ligation Reactions

This is a general protocol for use with the procedure for producing competent cells provided above. Please follow manufacturers’ instructions when using purchased competent cells.

1. Thaw a 100µl aliquot of competent cells on ice.
2. Transfer 100µl aliquot of the competent cells to a 17 × 100mm polypropylene tube prechilled on ice.
3. Add no more than 10ng of DNA in a maximum of 10µl from a ligation reaction to the cells and gently swirl the pipet tip to mix (do not mix by pipetting). Incubate on ice for 30 minutes.
4. Transfer the tubes from ice to a 42°C water bath and heat-shock for 45–60 seconds. Place on ice immediately to cool for 2 minutes.
5. Add 1ml of LB or SOC medium and incubate for 45 minutes at 37°C with shaking (~150rpm).
6. Plate 100–200µl of the transformation mix or an appropriate dilution, onto selection plates. If you suspect low ligation efficiency, take the remaining cells and pellet by a quick 10–20 second spin in a microcentrifuge. Pour off media and resuspend pellet in about 200µl of SOC and plate.

200cfu = 2 × 10⁵cfu/ng = 2 × 10⁸cfu/µg DNA

Transformation Controls

Controls help you figure out where things may have gone wrong with the cloning procedure. When transforming bacteria with your cloning reaction DNA, also determine transformation efficiency. Transforming a ligation control of cut, dephosphorylated vector without insert can tell you how many background colonies you can expect in your actual vector + insert ligation. 

Now that you’ve transformed your DNA and allowed the colonies to grow overnight, you need to determine if they contain the insert of interest. You can either screen them by colony PCR, or the more traditional plasmid miniprep followed by restriction digestion. Colony PCR is the most rapid initial screen. A plasmid miniprep will take an extra day to grow up the culture but will provide a lot of material for further analysis. Some researchers choose to do both but only perform minipreps on colonies  identified through colony PCR to contain the insert of interest.

Colony PCR

Colony PCR involves lysing the bacteria and amplifying a portion of the plasmid. If your cloning scheme will not maintain the orientation of the insert, you can even use colony PCR to screen for orientation. You can use either insert-specific primers or vector-specific primers to screen for recombinant plasmids. Simply combine a vector-specific primer with an insert-specific primer. 

PCR cloning using the A-overhangs left by Taq DNA Polymerase and an appropriately T-tailed vector (e.g., pGEM®-T Easy Vector) is not a technique that will retain orientation. The orientation can be rapidly assessed with colony PCR using vector-specific primers and insert-specific primers. For example, this technique was used for screening the orientation of a 1.8kb insert into the pGEM-T Easy Vector. Colony PCR was performed with the T7 Promoter Primer and either the insert-specific forward or reverse PCR primer. Eight white colonies were chosen from the cloning experiment for analysis. Clones with the T7 orientation will only produce a fragment with the T7 primer and reverse PCR primer, and clones in the opposite (SP6) orientation will only produce a fragment with the forward PCR primer as illustrated below.

Colony Prep for Colony PCR

1. Pick a well-isolated colony and transfer to 50µl of sterile water. Part of the colony may be transferred to LB medium containing the appropriate antibiotic for overnight culture and miniprep, if desired.
2. Boil the colony in sterile water for 10 minutes.
3. Centrifuge at 16,000 x g for 5 minutes.
4. Use 5µl of the supernatant in a 50µl PCR.

Screening by Plasmid Miniprep and Restriction Enzyme Digest

The classic method for screening colonies involves performing a plasmid miniprep followed by restriction digestion. Well-isolated colonies are picked from a plate and transferred to culture medium containing the appropriate antibiotic for selection. Proper sterile technique is important. Many different culture media formulations are commonly used for minipreps. Promega recommends LB medium supplemented with antibiotics for miniprep cultures. If a very rich medium like Terrific Broth is used, the bacteria can grow to very high cell densities and deplete the antibiotic. Once the antibiotic is depleted, the selection pressure to keep the plasmid is removed, and the plasmid may be lost.

You can inoculate the colony into 1–10ml of culture medium. If using a high-copy plasmid, 1–5ml (more typically, 1–2ml) is plenty. If you are using a low-copy plasmid, inoculate 10ml. Aerating the culture is very important for maximum cell density. A 17 × 100mm culture tube is fine for 1–2ml. If growing a larger volume, a 50ml sterile, disposable culture tube is better. Incubate the culture overnight (12–16 hours) with shaking (~250rpm). Remember, the greater the surface area, the greater the aeration. You can even grow miniprep cultures in sterile 25–50ml Erlenmeyer flasks.

Once the DNA is purified, a portion of the plasmid is screened by restriction digestion. For high-copy plasmids, you can obtain 4–10µg plasmid DNA per purification (1–5ml). For low-copy plasmids, you will obtain 1–3µg plasmid DNA per purification (10ml). Use 0.5–1µg of plasmid in your digest. Design the digest so that you can easily determine if your plasmid contains insert.
Note: Be sure to run uncut plasmid on the same gel for comparison.

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2. Higgins, N.P. and Cozzarelli, R. (1989) In: Recombinan DNA Methodology  Wu, R., Grossman, L. and Moldave, K., (Eds). Academic Press, Inc., San Diego, California.
3. Kaufman, D.L. and Evans, G.A. (1990) Restriction endonuclease cleavage at the termini of PCR products. BioTechniques 9, 304–6.
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5. Mole, S.E., Iggo, R.D. and Lane, D.P. (1989) Using the polymerase chain reaction to modify expression plasmids for epitope mapping. Nucl. Acids Res. 17, 3319.
6. Okazaki, R. et al. (1968) Proc. Natl. Acad. Sci. USA 59, 598.
7. Scharf, S.J., Horn, G.T. and Erlich, H.A. (1986) Direct cloning and sequence analysis of enzymatically amplified genomic sequences. Science 233, 1076–8.