Team:Paris Bettencourt/Protocols

Protocol 1: Heat Shock Transformation of E. coli

This protocol can be used to transform chemically competent (i.e. from CaCl2) with a miniprepped plasmid or a ligation product.

Note: Never vortex competent cells. Mix cells by gentle shaking.

1. Thaw competent cells on ice. These can be prepared using the CaCl2 protocol.
2. Place 20 ul of cells in a pre-chilled Eppendorf tube.
   • For an Intact Vector: Add 0.5 ul or less to the chilled cells
   • For a Ligation Product: Add 2-3 ul to the chilled cells.
3. Mix gently by flicking the tube.
4. Chill on ice for 10 minutes. This step is optional, but can improve yields when transforming a ligation product.
5. Heat shock at 42 °C for 30 seconds.
6. Return to ice for 2 minutes.
7. Add 200 ul LB medium and recover the cells by shaking at 37 °C.
   Another rich medium can substitute for the recovery.
   The recovery time varies with the antibiotic selection.
   Ampicillin: 15-30 minutes
   Kanamycin or Spectinomycin: 30-60 minutes
   Chloramphenicol: 60-120 minutes
8. Plate out the cells on selective LB.
   Use glass beads to spread the cells.
   The volume of cells plated depends on what is being transformed.
   
   • For an Intact Vector: High transformation efficiencies are expected. Plating out 10 ul of recovered cells should produce many colonies.
   • For a Ligation Product: Lower transformation efficiencies are expected. Therefore you can plate the entire 200 ul volume of recovered cells.
   Note: 200 ul is the maximum volume of liquid that an LB plate can absorb.
9. Incubate at 37 °C. Transformants should appear within 12 hrs.

Protocol 2: CaCl2 Competent Cells

This protocol makes 4 ml of competent cells, and can be easily scaled up to make more. The cells are typically stored in 110 ul aliquots, so this will make about 35 tubes. A typical transformation uses 20 ul of cells.

Note: Never vortex competent cells. Resuspend by pipetting with large Pasteur pipettes.

The night before:

1. The night before, inoculate a 5 ml culture and grow overnight with selection.

   The day of:

2. Dilute cells ~ 1:200 into selective media.
   For this example add 250 ul to 50 ml of selective media.
   Note: The protocol is easily scaled to increase the number of cells.
3. Grow the cells to an OD600 of 0.6 – 0.7.
   Use a large flask, 500ml, for good aeration.
   Use a baffled flask for fastest growth.
   This takes about 3 hours depending on the cells.
   Medium-heavy cloudiness by eye is fine.
4. Spin down the cells at 4 ºC, 4000 rpm, 15 minutes. Note: Keep the cells at 4 ºC from now on.
5. Resuspend cells in 15 ml, ice-cold 100 mM CaCl2. Leave on ice 4 hours to overnight.
6. Spin down the cells at 4 ºC, 4000 rpm, 15 minutes.
7. Resuspend cells in 4 ml, ice-cold 100 mM CaCl2 + 15% glycerol.
8. Aliquot into pre-chilled Eppendorf tubes. Use immediately or store at -80ºC.

Protocol 3: Glycerol Stocks

1. Pick Single colonies from agar plates
2. Innoculate 5ml LB broth overnight.
3. Add 750ml of overnight culture to 250ml of 60% glycerol in a cryotube.
4. Make two sets of Glycerol stocks freeze one at -20ºC and the other at -80ºC.

Protocol 4: Electroporation

Protocol: Electroporation (à la Datsenko and Wanner)

Preparation of Electrocompetent Cells
Note: Competent cells should never be vortexted, as this will cause them to lyse
and release salts into the media. Resuspend cells by pipeting up and down with a large
pasteur pipet. Once they are chilled, cells should be continuously cold.


1. The night before the transformation, start an overnight culture of cells.
   5 ml LB Amp.
   
   
2. The day of the transformation, dilute the cells 100X.
   100 ml LB Amp.
   Grow at 30°C for about 90 minutes.
   
   
3. Harvest the cells.
   When the cells reach an OD600 of between 0.6 and 0.8.
   Split the culture into 2x 50 ml falcon tubes, on ice.
   Centrifuge at 4 °C for 10 min at 4000 rpm.
   
   
4. Wash and combine the cells.
   Remove the supernatant.
   Resuspend the cells in 2x 25 ml of ice cold water.
   Combine the volumes in a single 50 ml falcon tube.
   
   
5. Wash the cells 2 more times.
   Centrifuge at 4 °C for 10 min at 4000 rpm.
   Resuspend in 50 ml of ice cold water.
   Repeat.
   
   
6. Wash and concentrate the cells for electroporation.
   Centrifuge at 4 °C for 10 min at 4000 rpm.
   Resuspend in 1-2 ml of ice cold water.
   We will use 200 ul of washed cells per transformation.
   

Dialysis of PCR or Digestion Products
Note: DNA for electroporation must be free of salts to avoid arcing.


1. Float a filter in a Petri dish filled with water.
   Millipore membrane filter 0.025 uM.
   
   
2. Pipet one drop of PCR product onto the filter.
   200 ng is needed per transformation.
   20 - 100 ul fits well on one filter.
   
   
3. Collect the drop after 30 - 45 minutes.
   The volume will change, but the DNA is not lost.

Protocol 5: Miniprep

Miniprep using the Thermo Scientific Miniprep Kit

1. Pellet 5ml of liquid culture (max speed, 1 min)
2. Discard supernatant
3. Resuspend the cells in 250ul of resuspension olution
4. Add 250ul of lysis solution, mix by inverting 4-6 times
5. Add 350ul of neutralization solution
6. Centrifuge for 5 min
7. Transfer supernatant to spin column
8. Centrifuge for 1 min
9. Discard flow through
10. Add 500 ul wash solution and centrifuge for 1 min , discard flow through(repeat this step)
11. Centrifuge for 1 min to remove left over liquid
12. Transfer the column on a 1.5ml tube
13. Add 50ul of elution buffer and incubate for 2 min
14. Centrifuge for 2 min
15. Nanodrop the concentration and freeze at -20°C

Protocol 6: PCR Purification

PCR purification using Thermo Scientific GeneJEt PCR Purification Kit

1. Add a 1:1 volume of Binding Buffer to completed PCR mixture (e.g. for every 100 uL of reaction mixture, add 100 uL of Binding Buffer). Mix thoroughly. Check the color of the solution. A yellow color indicates an optimal pH for DNA binding. If the color of the solution is orange or violet, add 10 uL of 3 M sodium acetate, pH 5.2 solution and mix. The color of the mix will become yellow.
   
   
2. Optional: if the DNA fragment is inf 500 bp, add a 1:2 volume of 100% isopropanol(e.g., 100 uL of isopropanol should be added to 100 uL of PCR mixture combined with 100 uL of Binding Buffer). Mix thoroughly.
   Note. If PCR mixture contains primer-dimers, purification without isopropanol is recommended. However, the yield of the target DNA fragment will be lower.
   
   
3. Transfer up to 800 uL of the solution from step 1(or optional step 2)to the GeneJET purification column. Centrifuge for 30-60 s. Discard the flow-through.
   Note. If the total volume exceeds 800 uL, the solution can be added to the column in stages. After the addition of 800 uL of solution, centrifuge the column for 30-60 s and discard flow-through. Repeat until the entire solution has been added to the column membrane.
   
   
4. Add 700 uL of Wash Buffer (diluted with the ethanol as described on p. 3) to the GeneJET purification column. Centrifuge for 30-60 s. Discard the flow-through and place the purification column back into the collection tube.
   
   
5. Centrifuge the empty GeneJET purification column for an additional 1 min to completely remove any residual wash buffer. Note.This step is essential as the presence of residual ethanol in the DNA sample may inhibit subsequent reactions
   
   
6. Transfer the GeneJET purification column to a clean 1.5 mL microcentrifuge tube (not included).Add 50 uL of Elution Buffer to the center of the GeneJET purification column membrane and centrifuge for 1 min.
   Note. For low DNA amounts the elution volumes can be reduced to increase DNA concentration. An elution volume between 20-50 uL does not significantly reduce the DNA yield. However, elution volumes less than 10 uL are not recommended. If DNA fragment is inf 10 kb, prewarm Elution Buffer to 65°C before applying to column. If the elution volume is 10 uL and DNA amount is inf 5 ug, incubate column for 1 min at room temperature before centrifugation.
   
   
7. Discard the GeneJET purification column and store the purified DNA at -20°C.