Colony PCR is a rapid screening technique used to determine whether bacterial colonies contain a plasmid with the correct DNA insert. Instead of purifying plasmid DNA from each colony first, the PCR is performed directly on a small amount of bacterial cells.
How Colony PCR Works
- Colony Selection
Individual bacterial colonies growing on an agar plate are picked using a sterile pipette tip or toothpick. Each colony is touched briefly, transferring a tiny amount of cells. The same tip is used to inoculate both the PCR tube and a master plate for later culture.
- Cell Lysis
The bacterial cells are heated to 95°C for several minutes during the initial denaturation step of the PCR. This heat treatment lyses the cells, releasing the plasmid DNA into the reaction mixture. The liberated DNA then serves as the template for amplification.
- PCR Amplification
Primers that flank the insert region in the plasmid are used. If the colony contains a plasmid with the correct insert, the PCR will produce a band at the expected size. If the plasmid is empty (no insert), the band will be smaller. If the colony does not contain the plasmid at all, no band will be produced.
- Gel Analysis
The PCR products are analyzed by agarose gel electrophoresis. Colonies that produce a band at the expected insert size are identified as positive. These colonies can then be grown for plasmid purification and further analysis.
- Advantages
Colony PCR is fast and inexpensive, allowing dozens of colonies to be screened in a few hours. It eliminates the need for plasmid purification before screening and is a standard first step in molecular cloning workflows.
Practical Colony PCR Protocol
Pick individual bacterial colonies from a selective agar plate using a sterile 10 µL pipette tip. Touch a single colony lightly — avoid transferring a large clump of cells. Streak the tip onto a fresh master plate (gridded LB-agar with antibiotic) to preserve the clone, then dip and swirl the same tip into a PCR tube containing 20–25 µL of master mix. For each colony, prepare the mix: 12.5 µL of 2× PCR master mix, 0.5 µL each of forward and reverse primers (10 µM), 1 µL of template (the colony), and water to 25 µL. Use vector-specific primers flanking the insert (e.g., M13 forward/reverse or T7/SP6) or insert-specific internal primers. Run the PCR with an extended initial denaturation at 95°C for 5–10 minutes to ensure complete bacterial lysis and DNA release. Follow by 30–35 cycles of 95°C for 30 s, 55°C for 30 s, and 72°C for 1 min per kb of expected insert, with a final extension at 72°C for 5 minutes. Analyze 5–10 µL of each reaction by agarose gel electrophoresis. A band at the expected insert size indicates a positive clone. If no band appears, the colony may lack the plasmid, contain an empty vector (producing a smaller band), or the lysis was insufficient — reduce the amount of bacterial cells transferred. For stubborn Gram-positive bacteria, add 0.5 µL of lysozyme (50 mg/mL) to the PCR mix or perform a freeze-thaw lysis prior to PCR.
Real-World Application
When cloning a 1.2 kb GFP expression cassette into pET-28a, transformants are plated on kanamycin agar. Of 24 colonies screened by colony PCR with T7 promoter and terminator primers, 18 show a 1.2 kb band, 4 show the empty vector band (300 bp), and 2 produce no band. The 18 positive clones are used for plasmid purification and sequencing, confirming successful cloning in under 4 hours.
