Preparation of plasmid minipreps

Our laboratory relies upon commercial plasmid purification kits which are highly reliable and produce very homogeneous plasmid preparations. Our protocols follow, and are adapted from the manufacturer’s recommendations. We currently prefer the use of the Zymo Research Zyppy Plasmid Miniprep kits.

Zymo Research Zippy Plasmid Miniprep Kit

• Centrifuge 1.6-2.0 mL¹ of cell culture at 14 krpm for 1 min, save pellet.
• Add 600 μL of water and resuspend pellet completely by vortexing.²
• Add 100 μL of 7X lysis buffer and mix by inversion 4-6 times. The solution should turn from opaque to clear blue indicating complete lysis.
• Within 2 minutes of lysis, add 350μL of cold neutralization buffer and mix by inversion 2-3 times. The solution should turn homogeneouly yellow if properly mixed and neutralized, and a yellowish precipitate of genomic DNA should form.
• Centrifuge at 14 krpm for 4 min.
• Carefully remove supernatant (about 900 μL) to spin columns equipped with catch tubes. Do not disturb the pellet.
• Centrifuge spin columns at 14 krpm for 15 s and discard flow-through.
• Wash with 200 μL of Endo-Wash buffer, centrifuge at 14 krpm for 15 s. (It is not necessary to empty the catch tube at this point)
• Wash with 400 μL of Zyppy Wash Buffer and centrifuge for 14 krpm for 30 s.
• Centrifuge at 14 krpm for 1 min to remove last traces of PE buffer from spin column
• Place spin column in a 1.5 mL microcentrifuge tube.³
• Add 30 μL of water directly to the column matrix, let stand 1 min, then centrifuge at 14 krpm for 15 s to collect DNA.⁴
• Use DNA immediately or store at –20 °C

Notes

1. It is possible, especially for low copy-number plasmids to collect cells from as much as 10 mL of overnight culture for this protocol. This can be accomplished by successive centrifugation of 1.6-2.0 mL aliquots of culture in the same microcentrifuge tube. The extra quantity of cells usually presents no problem in the plasmid purification protocol.
2. We have found that dragging the microcentrifuge tube rapidly back and forth across the holes in a microcentrifuge rack is highly effective in resuspending pelleted cells, and is usually superior to vortexing.
3. It may be necessary to decapitate the lids from the tubes in order to fit them into the microcentrifuge.
4. Note: Zyppy DNA minipreps are about 3X the concentration of Qiagen minipreps. Adjust quantities of DNA for sequencing or other downstream processing appropriately.

Qiagen miniprep spin kit

• Centrifuge 1.6-2.0 mL¹ of cell culture at 14 krpm for 3 min, save pellet.
• Add 250 μL of Buffer P1 and resuspend pellet completely by vortexing.²
• Add 250 μL of Buffer P2, mix gently by inversion, allow to lyse for no more than 5 min
• Add 350 μL of Buffer N3 to terminate lysis, mix gently by inversion
• Centrifuge at 14 krpm for 10 min, carefully remove supernatant to spin columns equipped with catch tubes
• Centrifuge spin columns at 6 krpm for 1 min, pulse at 14 krpm, and discard flow-through
• Wash with 500 μL of PB buffer, centrifuge at 6 krpm for 1 min, discard flow-through
• Wash with 750 μL of PE buffer, centrifuge at 6 krpm for 1 min, discard flow-through
• Centrifuge at 14 krpm for 1 min to remove last traces of PE buffer from spin column
• Place spin column in a 1.5 mL microcentrifuge tube.³
• Add 100 μL of water, let stand 1 min, then centrifuge at 6 krpm for 1 min, pulse at 14 krpm.
• Use DNA immediately or store at –20 °C

Notes

1. It is possible, especially for low copy-number plasmids to collect cells from as much as 10 mL of overnight culture for this protocol. This can be accomplished by successive centrifugation of 1.6-2.0 mL aliquots of culture in the same microcentrifuge tube. The extra quantity of cells usually presents no problem in the plasmid purification protocol.
2. We have found that dragging the microcentrifuge tube rapidly back and forth across the holes in a microcentrifuge rack is highly effective in resuspending pelleted cells, and is usually superior to vortexing.
3. It may be necessary to decapitate the lids from the tubes in order to fit them into the microcentrifuge.

Quantifying DNA

The quantity of DNA can be determined from its absorbance at 260 nm. Its quality can be estimated by measuring the A₂₆₀/A₂₈₀ ratio.

Pipet 5 μL of plasmid miniprep DNA (approx. 0.25 μg) into 195 μL of water in a 200 μL quartz silica cuvette. Measure the absorbance at 260 nm and at 280 nm.¹

• For ds-DNA, estimate the concentration using the formula A260 x 50 = μg/mL ds-DNA. Multiply by 40 (the dilution factor) to get the concentration of DNA in the original sample.²
• The A260/A280 ratio should be greater than 1.8 for pure ds-DNA. For most purposes, however, ds-DNA less pure than this can be used successfully for most molecular biology purposes.

Notes

1. Most diode array instruments will do this simultaneously.
2. For ss-DNA the formula is A₂₆₀ x 33 = μg/mL ss-DNA

Preparation of restriction digests

One of the most common operations in recombinant DNA methodology is the digestion of DNA with restriction enzymes. The following protocol is satisfactory for most applications.

• Mix the following reagents in a 1.5 mL microcentrifuge tube, adding the enzyme(s) last:
  • 1.5 μL of appropriate 10x buffer
  • 0.75 μL 20x acetylated BSA
  • 5-10 μL of DNA (approx. 0.5 μg)
  • 5-10 units of each restriction enzyme (< 1.5 μL total )
  • water to 15 μL
• Flick the tube gently to mix, spin briefly in a microcentrifuge to collect the content of the tube in the bottom, and incubate for 1-3 hr at 37 °C
• Dilute samples with 3 μL of 6x agarose gel loading buffer (see Molecular Biology Reagents for recipe), mix, and centrifuge to collect the sample in the bottom of the tube.
• Run out samples immediately in agarose gel electrophoresis to separate products

If using the FastDigest PstI kit, use the following quantities:

• 2 uL FastDigest 10X buffer
• 2-10 uL (up to 1 ug) DNA
• 1 uL FastDigest enzyme
• water to 20 uL
• Combine above reaction components at room temperature in the order indicated
• Mix gently and spin down
• Incubate at 37 °C for 5 min (plasmid and genomic DNA) or 30 min (PCR product)
• If FastDigest Green Buffer was used, sample is ready to be loaded in agarose gel immediately

Dephosphorylating vectors

When vector DNA is linearized by digestion by one or more restriction enzymes, it is often desirable to prevent self-ligation of the vector in subsequent cloning steps. This is absolutely required if a vector is digested with a restriction enzyme that leave blunt ds-DNA ends. The simplest way to prevent self-ligation is to remove the 5’-phosphate groups which are required for ligation by T4 DNA ligase. The protocol is conveniently carried out on a 15 μL restriction digestion mixture described above.

• Immediately after completion of restriction enzyme digestion, incubate the mixture at 65 °C for 20 min to inactivate restriction enzymes.
• Dilute the sample to 26 μL with water, add 3 μL 10x phosphatase buffer, and add 1 μL of calf intestinal alkaline phosphatase (1 unit/μL)
• Flick the tube gently to mix, spin briefly in a microcentrifuge to collect the content of the tube in the bottom, and incubate at 37 °C for 30 min
• Run out immediately in agarose gel electrophoresis to separate products

Phosphorylating oligonucleotides

When a DNA fragment is going to be ligated to a linearized, dephosphorylated vector with blunt ends, it is necessary for the DNA fragment to be 5’-phosphorylated for successful ligation with T4 DNA ligase. When the DNA fragment is a PCR product, this is most conveniently accomplished by phosphorylating the ss-DNA oligonucleotide primers used in PCR. A protocol for phosphorylating ss-DNA oligonucleotide primers is given below.

• Mix the following reagents in a 1.5 mL microcentrifuge tube, adding the enzyme(s) last:
  • 4 μL ss-DNA oligonucleotide (50 pmol/μL)
  • 5 μL 10x T4 ligase buffer with 10 mM ATP
  • 1 μL T4 polynucleotide kinase (10 u/μL)
  • water to 50 μL
• Flick the tube gently to mix, spin briefly in a microcentrifuge to collect the contents of the tube in the bottom,and incubate at 37 °C for 30 min
• Incubate at 70 °C for 10 min to inactivate enzymes
• Use solution immediately for PCR. Store unused portion at –20 °C

Ligation protocols

Standard ligation protocol

Ligation is a critical step in recombinant DNA methodology which accomplishes the “stitching together” of two disparate fragments of DNA. This is a common point of failure in cloning attempts. A robust protocol for ligating a PCR product to a linearized vector follows.

• Mix the following reagents in a 1.5 mL microcentrifuge tube, adding the enzyme(s) last:
  • 2-3 μL of linearized vector (approx. 50 ng)
  • 4-5 μL of PCR product
  • 1 μL 10x T4 ligase buffer with 10 mM ATP
  • 1-2 μL of T4 ligase (3-6 Weiss units per μL)
  • water to 10 μL
• Flick the tube gently to mix, spin briefly in a microcentrifuge to collect the contents of the tube in the bottom
• Incubate in the refrigerator (4 °C) overnight. This solution may be used directly to transform cells. Store unused portion at –20 °C.

Quick Ligation™ protocol

Our laboratory has experienced excellent ligation results using Quick T4 Ligase (New England Biolabs). In addition, this protocol takes substantially less time than traditional ligation, and is the preferred method whenever practical. The basic procedure follows.

• Mix the following reagents in a 1.5 mL microcentrifuge tube:
  • 1-3 μL of linearized vector (approx. 50 ng)
  • 4-5 μL of PCR product
  • water to 10 μL
• Add 10 μL of 2x Quick Ligation Buffer and mix gently
• Add 1 μL of Quick T4 Ligase and mix thoroughly
• Centrifuge briefly to collect the solution in the bottom of the microcentrifuge tube
• Incubate at room temperature for 5 minutes
• Chill on ice; use immediately for transformation or store at –20 ºC

Basic PCR protocol

PCR is used to amplify a particular DNA fragment which is flanked by sequences complementary to two flanking ss-DNA oligonucleotide primers. The following protocol is appropriate for Pfu Ultra, which does not need pre-annealing prior to adding polymerase:

• Mix the following reagents in a 0.5 mL PCR tube:
  • 1 μL of plasmid template (approx. 50 ng)
  • 0.5 μL of oligonucleotide primer #1 (50 pmol/μL)
  • 0.5 μL of oligonucleotide primer #2 (50 pmol/μL)
  • 2 μL dNTP mix (2.5 mM each)
  • 2.5 μL 10x polymerase buffer
  • water to 24.5 μL
• Flick the tube gently to mix, spin briefly in a microcentrifuge to collect the contents of the tube in the bottom
• Add 0.5 μL of Pfu Ultra(2.5 units/μL, Stratagene) (See note)
• Flick the tube gently to mix, spin briefly in a microcentrifuge to collect the contents of the tube in the bottom
• Perform PCR using the following segments:
  • Segment 1: 95°C for 2 min
  • Segment 2: 25-30 cycles of 95°C (30 s) → 62°C (30 s) → 72 °C (1 min or 1 min/kb of target). The annealing temperature (62°C) may be adjusted from 50-72°C as required to get product.
  • Segment 3: 72 °C (10 min) for extension, and then hold at 4 °C or place on ice.
• Dilute 20 μL of the PCR mix with 4 μL of 6x agarose gel loading buffer and run out on a 1% low-melt agarose gel
• Visualize bands on a UV transilluminator, cut out and purify desired PCR product (see Electrophoresis Protocols)

Note: It is important to add the Pfu Ultra last, as it has 3'-5' exonuclease activity, and will destroy the primers unless there are sufficient dNTPs present.

Touchdown PCR protocol

• Mix the following reagents in a 0.5 mL PCR tube:
  • 1 μL of plasmid template (approx. 50 ng)
  • 0.5 μL of oligonucleotide primer #1 (50 pmol/μL)
  • 0.5 μL of oligonucleotide primer #2 (50 pmol/μL)
  • 2 μL dNTP mix (2.5 mM each)
  • 2.5 μL 10x polymerase buffer
  • water to 24.5 μL
• Flick the tube gently to mix, spin briefly in a microcentrifuge to collect the contents of the tube in the bottom
• Add 0.5 μL of Pfu Ultra(2.5 units/μL, Stratagene) polymerase
• Perform 20 cycles of 95°C (30 s) → 65 °C (30 s) → 72 °C (1 min or 1min/kb of target); but instead of the middle temperature staying constant, it is lowered 0.5-1°C per cycle, with the final temp being 45-55°C.
• Perform an additional 20 cycles of 95°C (30 s) → 50 °C (30 s) → 72 °C (1 min or 1min/kb of target)
• At the end of cycling incubate for 10 min at 72 °C, and then hold at 4 °C or place on ice.
• Dilute 25 μL of the PCR mix with 5 μL of 6x agarose gel loading buffer and run out on a 1.5% low-melt agarose gel in two wells of 15μL
• Visualize bands on a UV transilluminator, cut out and purify desired PCR product (see Electrophoresis Protocols)

Designing and storing ss-DNA oligonucleotide primers

Commercially synthesized oligonucleotide primers are now quite inexpensive (typically 25¢ per nucleotide or less at scales appropriate for PCR) and can be ordered and received in a few days. We use primers from Integrated DNA Technologies, Coralville, IA. A number of design factors for successful PCR primers should be considered:

• PCR primers should be complementary to the DNA sequences flanking the 5’ and 3’ ends of the fragment to be copied; each primer should exactly match at least 18 nt of the fragment sequence
• Additional, non-complementary bases may be added to the 5’-end of either or both primers; this is commonly done to add unique restriction enzyme sequences to the PCR product to facilitate its cloning. There is generally no limit to how many “extra” bases can be added. For example, adding the sequence CCATGG to the 5’ end of a nucleotide primer would add a NcoI restriction enzyme site in the final product. If restriction enzyme sites are introduced into PCR products in this fashion it is strongly recommended that some extra bases be added to the 5’-end to facilitate the direct digestion of PCR products by restriction enzymes. The recommended additional sequence is “TGC.” So, for example, to add an NcoI site to a PCR product, the sequence TGCCCATGG should be added to the 5’-end of the complementary DNA sequence.
• PCR primers should normally have T_m values of 55-65 °C. T_m values can be estimated by the formula 4 x (G+C) + 2 x (A+T), or any number of web-based T_m calculators available on the internet can be utilized. Ideally, both PCR primers should have T_m values that are matched closely, but this is not a rigorous requirement.
• A typical PCR primer, with extra bases on the 5’-end for introducing unique restriction sites, is 24-28 nt long.
• Ideally, PCR primers should have approximately 50% GC content.
• Complementarity of the 3’-ends (especially the last three nt) for any pair of primers for PCR should be strictly avoided. Such self-complementary primers can anneal to each other and produce prodigious quantities of “primer-dimer” at the expense of the desired PCR product, especially if there is high GC content at the 3’ end of the primer. An example of PCR primers that can form “primer-dimers” is illustrated below:

• All PCR oligonucleotide pairs should be checked for possible primer-dimer formation prior to ordering or carrying out PCR.
• Reconstitute commercial primers in TE buffer at 500 pmol/μL and store at –80 deg C. Remove and dilute aliquots as required to 50 pmol/μL for routine PCR.

Site-directed mutagenesis by megaprimer PCR¹

Megaprimer PCR

This two-step PCR-based method is certainly one of the simplest and most efficient methods of introducing specific point mutations into a DNA fragment coding for a protein. The method is summarized in Figure 2 below:

Figure 2. PCR-based site-directed mutagenesis. Location of the introduced mutation is indicated by the star.

In the first round of PCR one flanking oligonucleotide primer (primer 1) is paired with an oligonucleotide primer (mutant primer) which is designed to introduce a point mutation in the desired location within the gene. The PCR product for this reaction (PCR-1) is gel purified and used as a primer in a second round of PCR with the other flanking primer (primer 2) to produce a DNA product corresponding to the entire gene, with the desired mutation included. The mutant primer should be designed in such a way as to have at least 9 exactly complementary nucleotides flanking the base mismatches required to introduce the desired mutation. Normally, the mutant codon should be checked against a usage table of codons for E. coli to ensure that the codon is not rarely used for protein translation.

Figure 3. Codon wheel. Read from the inside out for converting codons to amino acids.

The resulting product of the 2-step megaprimer PCR can be used for restriction digestion and ligation into a target overexpression plasmid, or can be used for whole-plasmid ligation-free cloning using MEGAWHOP PCR. It is possible to entirely skip the second PCR if doing MEGAWHOP cloining, the mutagenic fragment generated by the first PCR reaction can be purified and used directly in MEGAWHOP as described below.

A typical protocol for site-directed mutagenesis follows:

• Mix the following PCR mix in a 0.5 mL PCR tube:²
  • 1 μL of plasmid template (approx. 50 ng)
  • 0.5 μL of oligonucleotide primer #1 (50 pmol/μL)³
  • 0.5 μL of mutant primer (50 pmol/μL)
  • 2 μL dNTP mix (2.5 mM each)
  • 2.5 μL 10x polymerase buffer
  • water to 24.5 μL
• Perform PCR according the protocol previously described
• Dilute 25 μL of the PCR mixture with 5 μL of 6x agarose gel loading buffer.
• Load 15 μL of the sample into each of two lanes of a 1½% low-melt agarose gel and separate by electrophoresis; for PCR products, φX174/Hae III markers or a PCR ladder are necessary to locate the DNA fragment of the correct length, and should be loaded in a nearby lane
• Cut out the PCR product from the agarose gel, extract and purify using GeneClean Turbo.⁵
• For the second PCR reaction, mix in a 0.5 mL PCR tube:
  • 1 μL of plasmid template (approx. 50 ng)
  • 0.5 μL of oligonucleotide primer #2 (50 pmol/μL)
  • 10 μL of PCR product from first PCR⁴
  • 2 μL dNTP mix (2.5 mM each)
  • 2.5 μL 10x polymerase buffer
  • water to 24.5 μL
• Perform PCR according the protocol previously described
• Dilute 15 μL of the PCR mixture with 3 μL of 6x agarose gel loading buffer, and run out on a 1½% low-melt agarose gel and separate by electrophoresis, using φX174/Hae III markers or a PCR ladder in an adjacent lane to help identify the DNA fragment of the correct length.
• Cut out the PCR product from the agarose gel, extract and purify using GeneClean Turbo.⁵

Cloning into expression vector

• Prepare a restriction digestion using 5 μL of the concentrated, purified PCR product.
• Re-purify the digested DNA using GeneClean Turbo.
• Perform a ligation of 4-5 μL of a solution of this DNA fragment to the desired vector digested with the same enzymes (and perhaps dephosphorylated as well)
• Transform competent E. coli cells with this ligation mixture, spread on LB plates with the appropriate antibiotic, and grow up overnight at 37 °C
• Pick 2-3 colonies from this plate, grow up overnight in 5 mL of LB medium with the appropriate antibiotic, and prepare plasmid minipreps for each selected clone.
• Perform restriction digestions of miniprep plasmid DNA with the appropriate restriction endonucleases, and run out the products on a 1-1½% agarose gel to verify the plasmids contain the desired insert DNA.
• For clones showing evidence of the appropriate DNA insert in the vector, prepare frozen glycerol stocks from some of the overnight liquid culture. These cultures can be used to prepare samples of plasmid DNA for sequencing, or for conduction pilot overexpression trials (described later).

Notes

1. Sarkar, G. & Sommer, S. S. (1990) Biotechniques 8, 404-407
2. It is possible to perform this and the subsequent PCR protocol at half-scale, if desired.
3. Normally this primer contains extra nucleotides on the 5’ end to add a restriction enzyme site and facilitate direct restriction enzyme digestion of the eventual PCR product.
4. Use the entire purified product from the first PCR reaction.
5. DNA purification using GeneClean Turbo is described in Electrophoresis Protocols.

ite-directed mutagenesis by "Megaprimer Quick-Change" (MEGAWHOP) PCR

The advent of extremely high-fidelity DNA polymerases has made whole-plasmid PCR a practical method of introducing mutations into cloned gene products. The following method is a modification of the Stratagene QuikChange protocol that uses an easily synthesized megprimer PCR product as the mutagenic primer instead of the somewhat more problematic long oligonucleotides used in the standard QuikChange method, and is based on the method of Chen et al.¹. This is by far our preferred method for constructing site-directed variant plasmids.

• Construct the desired mutated gene or gene fragment using a one- or two-step megaprimer PCR method as described in the previous section. A one-step PCR to generate a mutation-containing megaprimer is sufficient to carry out MEGAWHOP. The megaprimer for MEGAWHOP should be > 100-150 bp to allow for efficient purification from the original PCR. (If the megaprimer is too short, poor recovery during purification may result in low primer concentration for MEGAWHOP.)
• Set up a whole-plasmid PCR using the following reagents:
  • 1 μL of plasmid template miniprep (approx. 50 ng)²
  • 5 μL of mutated gel-purified PCR product (approximately 100 ng)
  • 2 μL dNTP mix (2.5 mM each)
  • 2.5 μL 10x Pfu Ultra polymerase buffer
  • water to 24.5 μL
• Mix thoroughly and denature at 95°C for 2 min
• add 0.5 μL (1.25 U) Pfu Ultra polymerase
• Mix thoroughly and perform PCR as follows:
  • Perform 25-30 cycles of PCR:
    • 95°C for 1 min
    • 60°C for 1 min
    • 72°C for 1 min per kb of DNA product (e.g., 5 min for a 4000 bp plasmid + 1000 bp gene product)
  • Product extension at 72°C for 7 min
• Cool PCR mixture to 4°C
• Transfer PCR mixture to a 1.5 mL microcentrifuge tube and add 1 μL (10U) of DpnI to digest the original template plasmid DNA.
• Mix thoroughly and incubate at 37°C for 2-3 hours.
• Use 1-5 μL of this mixture to transform Z-competent cells as described in Microbiology Protocols. Spread 10-100 μL of transformed cells on pre-warmed LB-ampicillin plates.
• Clones that grow on LB-ampicillin plates must be screened by DNA sequencing to identify successfully mutated transformants.

Notes

1. Chen, G. J., et al. (2000) Biotechniques 28, 498-505.
2. The plasmid used must be from a dam+ host strain of E. coli so that that plasmid is methylated and will be digested by DpnI in the last step of this method. JM109 is a dam+ strain.

Polymerase Incomplete Primer Extension (PIPE) Cloning of Deletion Variants

PIPE¹ cloning is a ligation-independent method of mutagenesis that is especially relevant for the construction of deletion variants, e.g. N-terminal and C-terminal truncations of proteins. The method relies upon using two primers whose 5'-end are self-complementary over a range of 15-17 nucleotides. Under conditions where PCR extension of the template to double-stranded DNA is incomplete, the complementary single-stranded regions of the 5'-regions can self-anneal and circularize. When E. coli are transformed with this DNA, the nicked, circularized DNA is repaired (ligated) in vivo yielding competent plasmid.

Design of truncation primers

The following diagram shows an example of the design of a pair of primers that would result in a 4 amino acid N-terminal truncation of the target gene:

The double-stranded DNA sequence represents the expression plasmid (black) with the inserted target gene sequence (blue). The forward (truncation) primer is colored red, and the reverse primer, which can be used for any truncation variant, is colored green. Overlap region for self-annealing and in vivo ligation are in italics. Primers are typically designed with a 15 -17 nt overlap region and 18-21 nt complementary region.

PIPE PCR protocol

• Mix the following reagents in a 0.5 mL PCR tube:
  • 1 μL of 1:20miniprep plasmid template (approx. 5 ng)²
  • 0.5 μL of oligonucleotide primer #1 (50 pmol/μL)
  • 0.5 μL of oligonucleotide primer #2 (50 pmol/μL)
  • 2 μL dNTP mix (2.5 mM each)
  • 2.5 μL 10x polymerase buffer
  • water to 24.5 μL
• Flick the tube gently to mix, spin briefly in a microcentrifuge to collect the contents of the tube in the bottom
• Add 0.5 μL of Pfu Ultra(2.5 units/μL, Agilent)³
• Flick the tube gently to mix, spin briefly in a microcentrifuge to collect the contents of the tube in the bottom
• Perform PCR using the following segments:
  • Segment 1: 95°C for 2 min
  • Segment 2: 25-30 cycles of 95°C (30 s) → 55°C (45 s) → 68°C (14 min for typical whole plasmid amplificiation, approx. 5kbp). The annealing temperature (55°C) may be adjusted from 50-72°C as required to get product.
  • Segment 3: Hold at 4 °C or place on ice.
• Transfer PCR mixture to a 1.5 mL microcentrifuge tube and add 1 μL (10U) of DpnI to digest the original template plasmid DNA.²
• Mix thoroughly and incubate at 37°C for 1-3 hours.
• Use 1-5 μL of this mixture to transform Z-competent cells as described in Microbiology Protocols. Spread 10-100 μL of transformed cells on pre-warmed LB-ampicillin plates.
• Clones that grow on LB-ampicillin plates must be screened by DNA sequencing to identify successfully mutated transformants.

Notes

1. Klock, H. E., et al. (2008) Proteins, Struct. Func. Bioinfor. 71, 982-994.
2. The plasmid used must be from a dam+ host strain of E. coli so that that plasmid is methylated and will be digested by DpnI in the last step of this method. JM109 is a dam+ strain. The use of diluted template with DpnI digestion greatly reduces background transformants. The method is fairly tolerant of template concentrations of 0.2-10 ng.
3. It is important to add the Pfu Ultra last, as it has 3'-5' exonuclease activity, and will destroy the primers unless there are sufficient dNTPs present.

Enzyme-Free Cloning

Enzyme-free cloning is PCR-based ligation-free method based on the annealing of multiple PCR products of a vector and an insert that have overlapping adapter sequences [Tillett and Neilan (1999) ‘’Nucl. Acids Res.’’ 27, e26, “Enzyme-free cloning: a rapid method to clone PCR

products independent of vector restriction enzyme sites”]. This ligation-independent approach avoids often problematic double-enzyme restriction digests and ligation that often plagues the traditional methods of cloning. The method requires 8 oligonucleotide primers, a vector, and a DNA source for the gene insert. The primers are designed so that there is a 12-15 nt overlap region between the vector and insert PCR products. The following example shows how to design primers to clone a gene into a vector using this method. The diagram below shows the DNA sequence of the final assembled vector. DNA sequence belonging to the starting vector is in lower case; DNA sequence belonging to the insert is in upper case. The eight primers required to do the PCR reactions are overlaid above and below the sequence of the final, assembled plasmid. The primers have and overlap sequence of 12 nt and non-overlapping oligonucleotide sequence of 18 nt.

PCR protocol

Four PCR reactions are necessary to produce the proper fragments:

1. template = vector; primer VF-S and VR-L
2. template = vector; primer VF-L and VR-S
3. template = insert; primer PF-S and PR-L
4. template=insert; primer PF-L and PR-S

A basic PCR protocol using a high-fidelity DNA polymerase (e.g., Pfu) can be used to perform these reactions. A good starting point would be:

• 1 pg – 50 ng plasmid template; or 10-100 ng of insert ds-DNA
• 95°C for 30 s
• 50-60°C for 30 s
• 72°C for 1 min per 1000 nt
• 25-30 cycles
• Extension at 72°C for 7 min

If using pg quantities of plasmid template, DpnI digestion is not necessary. For ng quantities of plasmid template, DpnI digestion of template is desirable to reduce background transformation of empty plasmid. All four PCR products should be run out on a gel to verify success and separately purified using a spin column kit. (Each pair of plasmid PCR products and the insert PCR products could be combined, if desired, prior to purification from the gel slice.)

Annealing

If the each pair of PCR products (plasmid pair and insert pair) is mixed, denatured, and re-annealed, 25% of the annealed products will contain 5’ overhangs, and 25% will contain 3’ overhangs. The amplified insert overhangs will be complementary with the amplified vector overhangs and will allow direct association by mixing. (No ligation required.) The following is a suggested protocol.

• Equimolar quantities of the mixture of amplified vector and the mixture of amplified insert are mixed together in 50μL of hybridization buffer (10 mM Tris-Cl; pH 7.4, 100 mM NaCl; 1 mM EDTA, pH 8.0)
• Denaturation
  • Heat to 95°C for 3 min
• Annealing (4 cycles)
  • Cool to 65°C for 2 min
  • Cool to 25°C for 15 min

Transformation

Transform competent cells directly with 2-5 μL of the annealing mixture, and spread on an appropriate antibiotic plate. Colonies that grow on this plate should contain intact plasmid with the insert DNA.