Transforming E. coli with Engineered Plasmid



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Making Competent Cells (Inoue Method)

Inoue and colleagues developed this method in 1990. It works well for many strains commonly used in cloning. The original method calls for growing the overnight E. coli cultures at 18°C. We find that it still works well if they are grown overnight at 37°C. Alternatively, you can buy ready-to-use chemically competent cells, or electrocompetent cells, including BL21 (Catalog No. CMC0014), and SIG10 (Catalog No. CMC0001). Commercially prepared competent cells can be advantagous because their preparation has been optimized, providing specialized cells with high transformation efficiency for unique applications. See the Competent Cell Selection Guide for the complete list.

You will need:

DMSO (dimethyl sulfoxide) (Catalog No. D8418)
LB plates for streaking out the E. coli
3 x 250 ml SOB (Catalog No. H8032) for growing overnight cultures
Extra SOB for the starter culture
Shaking incubator
0.5 M PIPES (piperazine-1,2-bis[2-ethanesulfonic acid]) pH 6.7 (Catalog No. P8203)
Inoue transformation buffer
Sterile microfuge tubes
Sterile centrifuge tube with a capacity of at least 250 ml
Centrifuge capable of spinning such a tube with a force of 2,500 x g

Heat Shock Competent Cells Production Protocol

Day 1

  1. Pick a single E. coli colony from an LB plate that has been incubated overnight. Inoculate a starter culture of 2 ml of SOB medium in a 50 ml tube with the colony and incubate it for approximately 7 hours at 37oC with shaking.
  2. Prepare or thaw out an aliquot of the Inoue Transformation Buffer.
  3. Inoculate three flasks containing 250ml SOB medium with three different volumes of the starter culture to ensure that one of them reaches the desired OD the following morning. We suggest trying, 10 μl, 50 μl and 250 μl. Incubate the flasks overnight at 37°C with shaking.

Day 2

  1. The next morning read the OD600 of all three cultures. Continue to monitor the OD600.
  2. When the OD600 of one of the cultures reaches 0.55, transfer that culture to an iced water bath for 10 minutes. (Start chilling the microfuge tubes.)
  3. Harvest the cells by centrifugation for 10 min at 2,500 x g and 4°C.
  4. Pour away the supernatant and re-suspend the cells gently by pipetting in 80 ml ice-cold Inoue transformation buffer.
  5. Recover the cells by centrifugation for 10 min at 2,500 x g and 4°C.
  6. Pour away the supernatant and re-suspend the cells gently by pipetting in 10 ml ice-cold Inoue Transformation Buffer.
  7. Add 0.75 ml of DMSO, mix by swirling, and store the cells on ice for 10 minutes.
  8. Dispense 100 μl aliquots of cell suspension into chilled microfuge tubes.
  9. Snap-freeze the aliquots of competent cells in batches by dropping them into liquid nitrogen.

Store the aliquots in a -80°C freezer until needed.

Making Agar Plates

This recipe is designed to make approximately 50-60 plates of Luria-Bertani (LB) agar plates for growing E. Coli with standard plasmids. Alternatively, you can purchase pre-measured LB agar in an EZ-Mix™ format, Catalog No. L7533.

10g Tryptone (Catalog No. T2559)
5g Yeast extract (Catalog No. Y1625, Y1626)
10g NaCl (Catalog No. S3014)
15g Agar powder (Catalog No. L2897)
To 1 litre with water

Measure out the amounts above and add to a 1 litre bottle with a good working lid (See below). Fill with clean deionised water (ideally at least 18 megaohms). Tighten lid and shake to mix the liquid and powder, don’t expect to dissolve it all but simply to free the powder from the bottom and remove the major clumps. If not mixed properly the powder can bake on the bottom of the bottle. Undo the lid about half a turn; add some autoclave tape and then autoclave.

After autoclaving ensure that the lid is done up tight and do not allow to cool below 45-50 degrees. Agar generally sets at about 40 degrees. As a guide if you can hold the bottle comfortably in your hand then it is ready to pour and may actually be too cool already. Don’t add the antibiotic when the agar is too hot, this can affect the antibiotic stability and half life, particularly Ampicillin. Add the correct amount of antibiotic (see below) and mix by gentle swirling. Avoid getting bubbles as this will result in bubbly or uneven plates. Pour about 12-15ml per plate (Catalog No. Z617636). A simple method is to stack the plates up with the large sections (lids) upright. Lift the entire stack up using the lid of the bottom plate and using your other hand pour in the agar. Pour enough to fill the bottom and then a little bit more. Then put the stack back down and lift again with the next lid in the stack. Sounds inconsistent but with time this is very reproducible.

Alternatively, you can purchase prepared LB agar plates, with or without antibiotics:

LB Agar Plates (Catalog No. L5542)
LB Agar Plates with Ampicillin (Catalog No. L5667)
LB Agar Plates with Kanamycin (Catalog No. L0543)
LB Agar Plates with Carbenicillin (Catalog No. L0418)
LB Agar Plates with Tetracycline (Catalog No. L8795)

Tip 1: Many lab bottles have a blue or clear plastic rim which can go missing or get burnt by flaming. These blue rims prevent the agar running down the bottle when pouring the plates and keep an air tight seal after autoclaving. Make sure the rim is intact.

Tip 2: 1 litre is quite a lot of agar and antibiotic, scale down the recipe to suit your needs. I generally make 0.5 litres at a time ( about 30 Plates).

Antibiotic concentrations:

Prepare stock in water: add 1µl per 1ml of media
Kanamycin (Catalog No. K4000) – Stock 50mg/ml, final concentration 50µg/ml
Ampicillin (Catalog No. A9518) - Stock 100mg/ml, final concentration 100µg/ml
Streptomycin (Catalog No. S6501) – Stock 50mg/ml, final concentration 50µg/ml
Spectinomycin (Catalog No. S9007) - Stock 100mg/ml, final concentration 100µg/ml

Prepare stock in ethanol: Add 5µl per ml media
Chloramphenicol (Catalog No. C0378) - Stock 34mg/ml, final concentration 170µg/ml
Tetracycline HCL (Catalog No. 87128) - Stock 10mg/ml, final concentration 50µg/ml

Bacterial Transformation

The easiest ways to get DNA into bacteria are heat shock and electroporation. The principle of heat shock is exactly what it states, you have to shock the bacteria by heating. If it isn’t a shock then it doesn’t work, so keeping the bacteria cold until then is essential. This cannot be over emphasised. When the bacteria go from cold to hot it creates holes in the cell membranes and allows the DNA to enter the cell. Electroporation is a similar principle except the electricity makes the holes. These processes mimic, or are at least based on, the natural process of bacterial competence. Typically electroporation gives more colonies, but not always. For most DNA cloning applications heat shock works fine.

Bacterial Transformation Heat Shock Protocol (common method)

  1. Thaw one tube of your pre-made competent cells per DNA/ligation reaction or control reaction on ice and push the tube deep into the ice. Thawing takes about 5-10 minutes. Keep the cells as cold as possible and avoid touching the part of the tube containing the cells; a small amount of heat can significantly decrease the transformation process.
  2. Pre-chill 15ml Falcon Tubes (Catalog No. SIAL0791) on ice and transfer 3-4 μl of the ligation reaction (or control reaction) into each tube.
  3. Add 95 μl of competent cells into each ligation reaction and incubate on ice for 20 minutes (minimum). Longer is OK but we have only tested up to 45 minutes.
  4. Heat shock at 42°C for 90 seconds in either a heat block or water bath.
  5. Then add 1 ml of LB Broth (Catalog Nos. L3022, L2542, or L3522) or SOC medium (Catalog No. S1797) without antibiotic and incubate the cells in an incubation shaker at 37°C, 227RPM for 1 hour.
  6. Pour all the LB containing the transformed competent cells onto an agar plate containing the correct antibiotic.
  7. Leave the plate upright to dry with the lid slightly off in a class 1 hood or in a 37°C incubator for about 5-10 minutes. Do not do this in a hood that is used for mammalian tissue culture. Your colleagues will not be happy. Do not leave the plate to dry for too long as some of the bacteria may die.
  8. Incubate the plate overnight at 37°C. The colonies that will appear originate from single transformed cells and are resistant to the antibiotic due to the presence of the plasmid. Each colony will contain millions of identical copies of the same cell, hence the term clone.

Bacterial Transformation Heat Shock Protocol (alternative method)

  1. Thaw one tube of your pre-made competent cells per DNA/ligation reaction or control reaction on ice and push the tube deep into the ice. Don’t just place it on the ice or just in it; keep the tube as cold as you can. Thawing takes about 5-10 minutes. Avoid touching the part of the tube containing the cells, a small amount of heat can signficantly decrease the transformation process.
  2. Add 2-5 μl of each ligation reaction (or control reaction) to the 100μl of competent cells. As you pipette out the mixture stir the tip very quickly. Do not pipette up and down. The tip is probably at room temperature so remove it as quickly as possible.
  3. Incubate on ice for 15 minutes (minimum). Longer is OK but we have only tested up to 45 minutes.
  4. Heat shock at 37°C for 3 minutes in either a heat block or water bath.
  5. Then add 1ml of LB medium (Catalog Nos. L3522, L2542, or L3022) without antibiotic and incubate the cells for a further 15 minutes at 37°C.
  6. Pour all the LB containing the transformed competent cells onto an agar plate containing the correct antibiotic.
  7. Leave the plate upright to dry with the lid slightly off in a class 1 hood or in a 37°C incubator for about 5-10 minutes. Do not do this in a hood that is used for mammalian tissue culture. Your colleagues will not be happy. Replace the lid and turn the plate over when it is close to being dry. Do not leave the plate to dry for too long as some of the bacteria may die as the agar dries out, you will also end up with a very thin plate of agar!
  8. Incubate the plate overnight at 37°C. The colonies that will appear originate from single transformed cells and are resistant to the antibiotic due to the presence of the plasmid. Each colony will contain millions of identical copies of the same cell, hence the term clone.

Picking Bacterial Colonies

This step is easy and doesn't really warrant a separate protocol but given that it is an important part of the process we have included a brief description.

After you have performed a transformation with the appropriate controls you should be left with three bacterial plates. If everything has gone correctly you should have lots of colonies on the ligation plate, less colonies (or in a fantasy world no colonies) on the plate with no fragment but with ligase, and even less colonies (or again, no colonies) on the plates with no fragment and ligase. Often the background will be quite high on the control plates and this does not necessarily mean that the ligation hasn't worked as long as you have more colonies on the ligation plate. The number of colonies you pick for further screening is determined by the ratio of colonies from the control plate (that had ligase in the reaction) to the number of colonies on the ligation plate.

For example, if you have 10 times more colonies on the ligation plate compared to the control plate then in theory (if you are performing standard sticky ended directional cloning) 9 out of 10 of colonies you pick will have the correct fragment ligated into it (this doesn't normally actually work out to be true because of things like multimers or concatemers etc). In this case picking 5-10 colonies should be more than enough.

If you only have twice as many colonies on the ligation plate compared to the control, then picking 10 or more would be wise. The plate can be put in the fridge and more colonies can be picked if necessary. Don't go nuts when picking colonies, picking 50 is a waste of time unless you are doing a very difficult ligation.

It is important to acknowledge a failure if there is no difference between the plates. In this case, don't bother picking the colonies. Repeat the previous steps and try to decrease the background by either digesting for longer, dephosphorylating for longer, exposing to less UV, or try someone else's enzymes. Quite often an enzyme has just stopped working.

Picking Colonies Protocol

  1. After taking the plates out of the 37ºC incubator place them upside down (i.e. the way they were in the incubator) on the bench top.
  2. Using a pipette boy or similar instrument, pipette 3-5 ml of LB media (Catalog Nos. L3522, L2542, or L3022) containing the correct concentration of antibiotic into sterile 25 ml or 50 ml tubes (the number of tubes depends on how many you want to grow) or similar tube with a screw top and label them. The volume to volume ratio of the bacterial culture to the amount of air in the tube is important to allow the bacteria to grow to sufficient density, as an approximate guide never have less than a 1:3 ratio of liquid to air but ideally more.
  3. In one hand take a sterile pipette tip on the end of a pipette, with the other hand pick up the upside down plate containing the bacteria from the ligation. Turn the plate over in your hand so that the bacteria are now facing upwards towards you and touch the tip of the pipette tip gently to a bacterial colony that is completely isolated from any other colony.
  4. Now place the same tip with bacteria on it into one of the tubes containing LB media (from step 2) and move the tip around a bit to release some of the bacteria into the liquid. Some people simply eject the pipette tip into the media but if you do this you will need to recover it the next day.
  5. Culture the tubes overnight in an incubated orbital shaker at 37ºC at 190-225 rpm.

Note: The above procedure should be performed with sterile technique. Ideally in close proximity to a Bunsen burner, if this is not available a class 1 hood will suffice. In reality, it is probably more important not to cross contaminate each sample with each other because you are growing them all in the same antibiotic, than worrying about external bacteria contaminating your samples (because they shouldn't have the resistance cassette, unless they come from a previous experiment off your bench).

Freezing Bacterial Colonies

Bacterial colonies grown as liquid culture can be kept at 4°C for a few weeks without detriment.  Any clone that you have confirmed to contain your insert should be stored for longer-term access.  The most common method of clone storage is frozen glycerol stocks. For a room-temperature alternative (particularly useful for large samples numbers), consider the new technology used by Biomatrica’s CloneStable products (Catalog No. 93121-017).

Glycerol Stocks Protocol

  1. Prepare a 50% glycerol stock solution by mixing 5ml glycerol (Catalog No. G5516) with 5ml distilled, sterile water.  Mix well.
  2. For each colony being stored, mix 500 µl overnight culture with 500 µl 50% glycerol solution.  Mix well and dispense into a 2ml cryovial.
  3. Store at -80°C.