GenieClone Universal Mutagenesis Kit (MORV0033)

Components

Self-prepared materials: original plasmid, mutagenic primers, chemically competent cloning cells, ddH2O, PCR tubes and a thermal cycler.

Storage

Store at -30 ~ -15°C.

1. Design primers
   • Single-base (or continuous multi-base, within 50 bp) mutations: design one primer pair to amplify the whole plasmid by inverse PCR.
   • Discontinuous multi-site mutations: design one primer pair per site and amplify the plasmid in segments (e.g. AF + BR, BF + CR, CF + AR for three sites).
   • Primer structure: 5' - 15–21 bp reverse-complementary region + at least 15 bp non-complementary region - 3'.
   • Keep the GC content of the complementary region at 40%–60% and avoid repeats. A Tm >60°C from the mutation site to the primer 3' end is preferred (exclude bases 5' of the mutation from the calculation).
   • Place the mutation in the complementary region (introduce it on both primers) or in the non-complementary region of one primer. Do not place it at the primer end.
2. Amplify the target plasmid
   • Thaw and mix the PCR components thoroughly; return to -20°C promptly after use.

*Do not use dUTP, or uracil-containing primers or templates. Use as little template as gives normal amplification; ≤1 ng of freshly prepared plasmid is recommended.

*Use 5 sec/kb for fragments ≤10 kb and 10 sec/kb for fragments >10 kb. Check a small aliquot on an agarose gel before proceeding.

3. Digest the template with DpnI
   • Mix 1 μl DpnI (10 U/μl) with 40–50 μl amplification product, pipette to mix, spin down and incubate at 37°C for 15 min. If template input was >1 ng, extend digestion to 1 h.
   • If amplification is specific, use the digest directly for recombination (≤1/5 of the recombination volume). Otherwise, gel-purify the target product after digestion.
   • Absorbance-based concentration readings are reliable only for purified product with A260/A280 of 1.8–2.0.
4. Calculate DNA input
   • Single-site and multi-site mutagenesis: 0.03 pmol of each fragment = [0.02 × fragment bp] ng.
   • Discontinuous double-site mutagenesis: use a 1:2 molar ratio, 0.03 pmol of the longer fragment ([0.02 × bp] ng) and 0.06 pmol of the shorter fragment ([0.04 × bp] ng). Example: AB 1 kb = 40 ng; BA 5 kb = 100 ng.
   • Keep each input within 10–100 ng; use the nearest limit if the calculated value falls outside this range. Unpurified digest must not exceed 2 μl in total.
5. Set up the recombination reaction on ice

^a A negative control is recommended for multi-site reactions, as excess template or incomplete DpnI digestion can give a high false-positive background.
^b Amount of each fragment calculated as in step 4.

• Mix gently by pipetting (do not vortex) and spin down.
• Incubate at 50°C for 5 min (single-site or discontinuous double-site) or 15 min (discontinuous multi-site), then chill immediately at 4°C or on ice. A thermal cycler is recommended for accurate temperature control.
• The recombinant product can be stored at -20°C for one week.

6. Transform
   • Thaw cloning competent cells on ice. Add 5–10 μl recombination product to 100 μl cells (≤1/10 of the cell volume), flick to mix (do not vortex) and keep on ice for 30 min.
   • Heat-shock at 42°C for 30 sec, then place on ice for 2–3 min.
   • Add 900 μl SOC or LB medium (no antibiotic) and shake at 37°C, 200–250 rpm for 1 h.
   • Centrifuge at 5,000 rpm (2,400 × g) for 5 min, discard 900 μl supernatant, resuspend the cells in the remaining medium and spread on a pre-warmed selective LB plate.
   • Incubate at 37°C for 12–16 h.
7. Identify clones
   • If the plate has clearly more colonies than the negative control, pick several, grow overnight in selective LB medium, extract plasmids and confirm by Sanger sequencing.

Q1: What should I do if the plasmid template does not amplify?

• Check the primer design against the guidance; repeat the experiment in case the reaction was set up incorrectly; adjust the amplification program; and use freshly prepared plasmid, as long-term storage and repeated freeze–thaw can break, nick or degrade the template.

Q2: Why are there few or no colonies on the plate?

• Check that primers include 15–20 bp homology arms with 40%–60% GC; use the recommended DNA amounts and ratios; keep unpurified DNA ≤2 μl (1/5 of the reaction), or gel-purify and dissolve in ddH₂O; and use cloning competent cells (e.g. DH5α or XL10, not expression strains) with transformation efficiency >10⁸ cfu/μg. Keep the transformation volume ≤1/10 of the cell volume.

Q3: Why was the mutation not introduced correctly?

• Possible causes are incorrect primer design; an unmethylated template (DpnI only recognises methylated DNA, so prepare plasmid from a host without methylase defects); or too much template, which leads to incomplete DpnI digestion. For most plasmids, 1 ng of template is sufficient.

Q4: Why are there mutations at non-target sites?

• The template may carry unknown mutations, so verify it by sequencing. Too many cycles can also introduce errors: use ≤35 cycles, and ≤30 if amplification is efficient.

Q5: How can I check competent cell efficiency?

• Transform 0.1 ng of plasmid and plate 1/10 of the culture. About 1,000 colonies corresponds to a transformation efficiency of 10⁸ cfu/μg.