Transformation¶
Bench cheatsheet
KCM Heat-Shock Transformation — the one-page version of this protocol. It is the same as the printed copies at the bench.
After assembly, the circular plasmid has been generated, but it is mixed with other DNAs—some incomplete or mutant. By passing the material through cells, we isolate a single one of these sequences and amplify it a billion-fold.
Transformation is the process in which a bacterium takes up DNA from the environment. While some bacteria (like B. subtilis) do this naturally, E. coli requires preparation to become “competent.”
There are two common methods:
🔌 Electroporation¶
Electroporation involves preparing cells in salt-free, ice-cold water or 10% glycerol and shocking them with a brief electric pulse in a special cuvette. This creates pores in the membrane, allowing DNA to enter. It is:
- ~100× more efficient than heat shock
- More expensive (cuvettes are single-use)
- Sensitive to salt contamination
- Ideal for library transformations needing high colony counts
🌡️ Heat Shock (used in pP6)¶
We use the TSS method for heat shock, where cells are suspended in a PEG-salt mix. You add DNA and a small amount of KCM buffer, chill, then heat shock at 42°C for 90 seconds.
📄 How the two solutions are made: TSS and 5× KCM. The method is from Chung, Niemela & Miller, PNAS 1989.
TSS cells can be frozen in aliquots and remain competent for years. TSS cells are easier to work with during transformation and are well-suited for routine cloning workflows.
In dilute conditions, as in pP6, most cells take up just one plasmid, resulting in unique colonies.
Rescue Step?¶
Some antibiotics (e.g., kanamycin, chloramphenicol) block translation immediately. To survive, cells must first express the resistance gene—this requires a rescue step (1 hour outgrowth in rich media before plating).
Ampicillin, used in pP6, is different: it kills only actively growing cells. Cells recover naturally after transformation, so no rescue is required.
Amp vs. Carbenicillin¶
Both are β-lactam antibiotics. The bla gene on pP6 confers resistance to both.
- Ampicillin breaks down when heated (e.g., during plate pouring)
- Carbenicillin is more stable, so we use Carb in practice
Cells without plasmid die at 5 µg/mL; with plasmid, they survive over 1000 µg/mL—a wide window for selection.
Transformation Efficiency¶
Transformation efficiency measures how effectively DNA is taken up and expressed by cells. It's typically calculated as:
CFU/µg DNA = (Colonies on plate) ÷ (amount of plasmid DNA in µg)
Our heat shock competent cells typically yield around 10⁶ CFU/µg DNA. With electroporation, efficiencies as high as 10¹¹ CFU/µg DNA can be achieved.
This varies based on:
- Method (electroporation ≫ heat shock)
- Competency of the cells
- DNA purity and volume
- Handling technique
Don’t worry about calculating it for pP6, but understand that low DNA input typically gives 1 DNA per cell, which is ideal when selecting single clones.
Handling Competent Cells¶
Competent cells are fragile. To maintain their viability:
- Keep them cold at all times (on ice or in a cold block).
- Do not pipette vigorously. Stir gently with the tip, then pipette up and down once or twice carefully.
- Avoid bubbles—they indicate lysis.
- Aliquot immediately after prep—cells degrade with freeze-thaw cycles.
- Do not re-freeze thawed competent cells—even one additional freeze-thaw cycle can dramatically reduce their transformation efficiency.
Note: You will not make your own competent cells. We prepare large batches once a year using the TSS method and store them at -80°C. Your instructor or supervisor will provide a frozen aliquot when needed.
Plate Hygiene and Freshness¶
A petri dish is not a shelf-stable reagent. Before you plate, check that the plate you are about to use is still good. A bad plate costs you a day and can give you colonies that do not carry your plasmid.
How long fresh plates keep. Plates are stored sealed, upside-down, at 4 °C, in a bag labeled with the antibiotic and the pour date.
- Carbenicillin plates: use within 4 weeks of the pour date. Throw them out at 6 weeks no matter how they look.
- Ampicillin plates: use within 2 weeks. Ampicillin breaks down faster than carbenicillin.
- A bag with no pour date on it is treated as expired.
Why that number: standard cloning practice puts β-lactam (amp/carb) plates at about one month at 4 °C. Carbenicillin is more stable than ampicillin, so it gets the longer window.
Contamination spreads through a bag.
- If one plate in a bag is contaminated, the others are at risk. Mold spores and bacteria move between plates in the bag, and they keep growing slowly in the fridge. The longer a bag sits, the more contaminated it gets.
- If you see any growth on an unused plate, treat the whole bag as suspect. Don't use its plates, and tell whoever pours plates.
- Take out only the plates you need and close the bag right away. Do not put unused plates back once they have been at the bench.
- Keep plates upside-down so condensation drips onto the lid, not onto the agar.
Plates dry out.
- Plates must be wrapped in saran wrap, sealed with parafilm, or kept in a closed bag. Even then they slowly dry out. Sealing slows drying; it does not stop it.
- Signs a plate is too dry: the agar pulls away from the side of the dish, shrinks or cracks, or turns a darker brown. Cells grow poorly on it, colonies come up small, and the antibiotic is more concentrated than intended. Throw it out.
Antibiotics are unstable.
- Ampicillin and carbenicillin break down over time, and faster when warm. A plate left out on the bench ages quickly — put plates back in the fridge.
- What an old plate looks like: lots of satellite colonies (small colonies crowded around a real one, growing where the antibiotic has been used up), or growth on a plate that should have none. On a plate with weakened antibiotic, colonies without your plasmid can survive.
After your plates have grown.
- Grown plates go in the culture fridge, not back with the fresh plates. Wrap them in parafilm and store them upside-down.
- They still dry out, and the cells on them die. Expect 1–2 weeks even when wrapped.
- A grown plate is a record, not storage. To keep a clone, restreak it onto a fresh plate or make a glycerol stock at −80 °C.
Plating¶
Plating is done with sterile glass beads at one of the lab's two sterile benches.
1. Prepare the bench.
Spray the working surface with 70% ethanol and wipe it down with paper towels before you
begin.
2. Label the plate.
Write your label, the date, and your name on the bottom of the plate before you put
anything on it. See Labeling and Inventory below.
3. Add the beads.
Pour sterile glass beads onto the surface of the plate — about 10 is typical, and the
exact number doesn't matter much. Sterile beads are kept in glass bottles marked with
autoclave tape.
4. Add the cells.
Pipette 50–200 µL of your cell suspension onto the plate.
5. Spread.
Cap the plate and shake it side to side — a straight back-and-forth motion. Watch the
beads to confirm they are travelling across the whole surface.
Do not swirl. A circular motion just drives the liquid out to the perimeter of the plate and leaves the middle unspread. Back and forth, not around.
6. Recover the beads.
Uncap the plate and pour the used beads through a funnel into the collection beaker.
The beads are not trash. They are collected, washed, and re-sterilized for reuse. Do not throw them away.
7. Incubate.
Recap the plate, invert it (lid on the bottom), and place it in the incubator — not
a shaker.
No open flames at the sterile bench. Flame sterilization and flamed metal spreaders are a supervisor-only technique carried out elsewhere in the lab, and are not part of this procedure. For plating, you use glass beads and 70% ethanol.
Labeling and Inventory¶
Be consistent and thorough:
- Write label, date, and your name on the plate bottom (not the lid since those can fall off).
- Match plate name to your sample ID and lab sheet entry (e.g.,
pP6-79). - Return any used DNA tubes to their storage rack and note usage if needed.
Controls¶
If you select with anything other than carbenicillin, run the restreak, positive and negative controls beside your plates, as your lab sheet lists them. Without them, a blank plate can't tell you whether the DNA, the cells or the plates failed. See Control Strains and Plasmids.