iGEM 2019 Bacterial Transformation Protocol
Introduction
Make sure you have competent cells (you can use this protocol to test them). It is important to have efficient competent cells because transformations performed with ligation products usually do not yield as many colonies due to the low DNA concentration in the ligation mixture. This means that you may see different results doing this test than you will at the end of the 3A Assembly protocol, or any other ligation.
This protocol for heat-shock transformation can be used on chemically-competent E. coli cells.
Materials
- 70% ethanol
- Paper towels
- Lab marker / Sharpie
- 1.5 mL microcentrifuge tubes
- Container for ice
- Ice
- Competent cell aliquot(s)
- Positive Control (Competent Cell Test Kit has this)
- Agar plates with appropriate antibiotic (CAM for competent test kit)
- 42°C Water bath (or hot water source and thermometer)
- 37°C Incubators (oven and shaker)
- SOC media
- Sterile glass beads or sterile cell spreader
- Pipettor
- Pipette tips
Protocol estimated time: 30 minutes active, 1.5 hours incubation
1. Clean your working area by wiping it down with 70% ethanol.
2. Thaw competent cells on ice. Label one 1.5 mL microcentrifuge tube for each transformation and then pre-chill by placing the tubes on ice.
Do triplicates (3 each) of each concentration if possible, so you can calculate an average colony yield.
3. Spin down the DNA tubes from your positive control (or Competency Test Kit) to collect all of the DNA into the bottom of each tube prior to use. A quick spin of 20-30 seconds at 8,000-10,000 rpm will be sufficient. Note: There should be 50 µL of DNA in each tube sent in the Kit.
4. Pipet 1 µL of DNA into each microcentrifuge tube. Note concentrations.
5. Pipet 50 µL of competent cells into each tube. Flick the tube gently with your finger to mix.
6. Incubate on ice for 30 minutes.
Pre-heat water bath now to 42°C. Otherwise, hot water and an accurate thermometer works, too!
7. Heat-shock the cells by placing them into the water bath for 45 seconds (no longer than 1 min). Be careful to keep the lids of the tubes above the water level, and keep the ice close by.
8. Immediately transfer the tubes back to ice, and incubate on ice for 5 minutes.
9. Add 950 µL of SOC media per tube, and incubate at 37°C for 1 hour shaking at 200-300rpm.
Prepare the agar plates during this time: label them, and add sterile glass beads if using beads to spread the mixture.
10. Pipet 100 µL from each tube onto the appropriate plate, and spread the mixture evenly across the plate. Incubate at 37°C overnight or approximately 16 hours. Position the plates with the agar side at the top, and the lid at the bottom.
To test for Competency: Count the number of colonies on a light field or a dark background, such as a lab bench. Use the following equation to calculate your competent cell efficiency. If you've done triplicates of each sample, use the average cell colony count in the calculation.
- Efficiency (in cfu/µg) = [colonies on plate (cfu) / Amount of DNA plated (ng)] x 1000 (ng/µg)
- Note: The measurement "Amount of DNA plated" refers to how much DNA was plated onto each agar plate, not the total amount of DNA used per transformation. You can calculate this number using the following equation: Amount of DNA plated (ng) = Volume DNA added (1 µL) x concentration of DNA (refer to the vial, convert to ng/µL) x [volume plated (100 µL) / total reaction volume (1000 µL)]
‘Addgene’ Inoculating an Overnight Liquid Culture
1. Prepare liquid LB. For example, to make 400 mL of LB, weigh out the following into a 500 mL glass bottle:
- 4g NaCl
- 4g Tryptone
- 2 g Yeast Extract
- dH2O to 400 mL
Note: If your lab has pre-mixed LB agar powder, use the suggested amount, instead of the other dry ingredients above.
Loosely close the cap on the bottle (do NOT close all the way or the bottle may explode!) and then loosely cover the entire top of the bottle with aluminum foil. Autoclave and allow to cool to room temperature. Now screw on the top of the bottle and store the LB at room temperature.
2. When ready to grow your culture, add liquid LB to a tube or flask and add the appropriate antibiotic to the correct concentration (see table below).
Note: If you intend to do a mini-prep you will usually want to start 2 mL in a falcon tube, but for larger preps you might want to use as much as a liter of LB in a 2 L Erlenmeyer flask.
3. Using a sterile pipette tip or toothpick, select a single colony from your LB agar plate.
4. Drop the tip or toothpick into the liquid LB + antibiotic and swirl.
5. Loosely cover the culture with sterile aluminum foil or a cap that is not air tight.
6.Incubate bacterial culture at 37°C for 12-18 hr in a shaking incubator.
Note: Some plasmids or strains require growth at 30°C. If so, you will likely need to grow for a longer time to get the correct density of bacteria since they will grow more slowly at lower temperatures.
7. After incubation, check for growth, which is characterized by a cloudy haze in the media (see right).
Note: Some protocols require bacteria to be in the log phase of growth. Check the instructions for your specific protocol and conduct an OD600 to measure the density of your culture if needed.
Note: A good negative control is LB media + antibiotic without any bacteria inoculated. You should see no growth in this culture after overnight incubation.
‘Addgene’ Glycerol Stock Preparation Protocol
1. Follow the steps for Inoculating an Overnight Liquid Culture
2. After you have bacterial growth, add 500 μL of the overnight culture to 500 μL of 50% glycerol in a 2 mL screw top tube or cryovial and gently mix.
Note: Make the 50% glycerol solution by diluting 100% glycerol in dH20.
Note: Snap top tubes are not recommended as they can open unexpectedly at -80°C.
3. Freeze the glycerol stock tube at -80°C. The stock is now stable for years, as long as it is kept at -80°C. Subsequent freeze and thaw cycles reduce shelf life.
4. To recover bacteria from your glycerol stock, open the tube and use a sterile loop, toothpick, or pipette tip to scrape some of the frozen bacteria off of the top. Do not let the glycerol stock unthaw! Streak the bacteria onto an LB agar plate.
5. Grow your bacteria overnight at the appropriate temperature. Growth conditions, including copy number and growth temperature, can be found on your plasmid's information page. The next day you will be able to start an overnight culture for plasmid DNA prep the following day.
Zymogen Mini Prep
Day 1
1. Inoculate cultures with bacteria transformed with the plasmid of interest the day before your miniprep.
Optimal volume ratio of culture to tube for aeration is 1:5
Day 2
2. Make sure all reagents and buffers* are available and at the proper temperatures before you begin.
Check that P2 and binding buffers have not precipitated, incubate 10min at 30-37 ̊C if necessary.
3. Centrifuge 0.5-5 ml1 of bacterial culture in a clear 1.5 ml tube at full speed for 15- 20 seconds in a microcentrifuge. Discard supernatant.
4. Add 250 μl of ZymoPURETM P1 (Red) to the bacterial cell pellet and resuspend completely by vortexing or pipetting.
5. Add 250 μl of ZymoPURETM P2 (Green) and immediately mix by gently inverting the tube 6-8 times. Do not vortex! Let sit at room temperature for 2-3 minutes. Cells are completely lysed when the solution appears clear, purple, and viscous.
Do NOT allow lysis to occur for longer than 3min as this may damage the DNA
6. Add 250 μl of ice-cold ZymoPURETM P3 (Yellow) and mix thoroughly by inversion. Do not vortex! Invert the tube an additional 3-4 times after the sample turns completely yellow. The sample will turn pale yellow when the neutralization is complete and a yellowish precipitate will form.
7. Incubate the neutralized lysate on ice for 5 minutes.
8. Centrifuge the neutralized lysate for 5 minutes at 16,000 x g.
9. Transfer 600 μl of supernatant from step 6 into a clean 1.5 ml microcentrifuge tube. Be careful not to disturb the yellow pellet and avoid transferring any cellular debris to the new tube.
Less than 600μl will affect concentration for binding, but make sure not to contaminate with protein precipitate...
10. Add 275 μl of ZymoPURETM Binding Buffer to the cleared lysate from step 7 and mix thoroughly by inverting the capped tube 8 times.
11. Place a Zymo-SpinTM II-P Column in a Collection Tube and transfer the entire mixture from step 8 into the Zymo-SpinTM II-P Column. Do NOT spin yet.
12.Incubate the Zymo-SpinTM II-P/Collection Tube assembly at room temperature for 2 minutes and then centrifuge at 5,000 x g for 1 min. Discard the flow-through.
The total capacity of the collection tube is 800μL so discard flow through any time it’s getting close to contaminating the bottom of the column
13. Add 800 μl of ZymoPURETM Wash 1 to the Zymo-SpinTM II-P Column and centrifuge at 5,000 x g for 1 min. Discard the flow through.
14. Add 800 μl of ZymoPURETM Wash 2 to the Zymo-SpinTM II-P Column and centrifuge at 5,000 x g for 1 min. Discard the flow through.
15. Add 200 μl of ZymoPURETM Wash 2 to the Zymo-SpinTM II-P Column and centrifuge at 5,000 x g for 1 min. Discard the flow through.
16. Centrifuge the Zymo-SpinTM II-P Column at ≥ 10,000 x g for 1 minute in order to remove any residual wash buffer.
17. Transfer the Zymo-SpinTM II-P Column into a clean 1.5 ml tube and add 25 μl of ZymoPURETM Elution Buffer* directly to the column matrix. Incubate at room temperature for 2 minutes, and then centrifuge at ≥ 10,000 x g for 1 minute in a microcentrifuge. Store the eluted plasmid DNA at ≤ -20°C.
Pre-warm Elution buffer to 50 ̊C and/or increase incubation period to 5min
GreenGel Gel Staining Protocol
Staining Protocols
Because nucleic acid binding dyes can affect DNA migration during electrophoresis, post-staining of gels is highly recommended. Post-staining with GelGreen® results in superior sensitivity and eliminates the possibility of dye interference with DNA migration. Post-staining with GelGreen® is simple, requiring no destaining and no special buffer. GelGreen® also can be included in agarose gels using the precast method. While the precast protocol is more convenient, some DNA samples may experience migration retardation or compromised resolution in the presence of GelGreen®. Thus, the poststaining and precast protocols should be compared to determine which one better meets your needs. Although GelGreen® has undergone extensive safety testing, Biotium recommends following universal safety precautions when working in the laboratory.
Post-staining Protocol 1
1. Run gels as usual according to your standard protocol.
2. Dilute the GelGreen® 10,000X stock reagent ~3,300 fold to make a 3X staining solution in H2 O. Note: including 0.1 M NaCl in the staining solution enhances sensitivity, but may promote dye precipitation if the gel stain is reused.
3. Carefully place the gel in a suitable polypropylene container. Gently add a sufficient amount of the 3X staining solution to submerge the gel.
4. Agitate the gel gently at room temperature for ~30 minutes.
5. Image the stained gel with a 254 nm transilluminator, a Dark Reader® or a similar transilluminator, or a laser-based gel scanner using a long path green filter such as a SYBR® filter or GelStar® filter.
6. Staining solution can be reused at least 2-3 times. Store staining solution at room temperature protected from light.
Precast Protocol
Note: The precast protocol is not recommended for polyacrylamide gels. Use the post-staining protocol for acrylamide gels.
1. Prepare molten agarose gel solution using your standard protocol.
2. Dilute the GelGreen® 10,000X stock reagent into the molten agarose gel solution at 1:10,000 and mix thoroughly. GelGreen® can be added while the solution is still hot.
3. Cast the gel and allow it to solidify. Load samples and run the gels using your standard protocol.
4. Image the stained gel with a 254 nm transilluminator, a Dark Reader® or a similar transilluminator, or a laser-based gel scanner using a long path green filter such as a SYBR® filter or GelStar® filter.
Gel Electrophoresis Protocol
We used the highlighted range below:
1. Measure out the appropriate mass of agarose into an Erlenmeyer flask and then add the appropriate volume of TE buffer using the chart. The volume required depends on your gelbox / casting system -- 50mL makes a good, thick gel for a 7x10cm gelbox.
2. Microwave until the agarose is fully melted. This depends strongly on your microwave, but 90 seconds at full power or 3 minutes at half power seem to provide decent results. Make sure to not burn the agarose and that nothing bubbles over.
3. At this point add your DNA stain - if using the Pre-Cast method
Dilute the GelGreen™10,000X stock reagent into the molten agarose gel solution at 1:10,000 and mix thoroughly. GelGreen™ can be added while the gel solution is still hot. If you are making a 50ml solution, this would be 5µl.
4.Let the agarose cool on your bench until touching the bottom of the beaker with your bare hand doesn't burn you (~5 minutes for a 50mL gel).
5. While waiting to cool, seal the open edges of your gel box with one long piece of masking tape on each side. Make sure it is sealed well or the gel will leak.
6. Insert a comb into the gelbox.
7. Pour the agarose solution into the taped gelbox. Carefully pop or shove to the side any bubbles, and let it cool for about 30 minutes until the gel is solid.
8. Carefully pull the comb out of the gelbox.
9. Carefully slide the solidified gel into the electrophoresis chamber. Make sure that the wells are on the same side as the anodes.
10. Fill the box with running buffer until the well is submerged.
11. Prepare the DNA constructs:
- With a micropipette, add 5 microliters of each DNA construct into a separate centrifuge tube.
- With a micropipette, add 2 microliters of gel electrophoresis dye into each centrifuge tube. Make sure to aim the dye at the DNA in the tubes and ‘mix’ the solution through repetitive pipetting.
12. On the first well, load 5 microliters of 100bp DNA.
13. On the second well load 4 microliters of Lambda HindIII DNA.
14. Load the rest of the wells with 5 microliters of constructs each.
15. Connect the anodes and the cathodes and turn on the power supply. Start from medium.
16. Run the gel for 30 to 60 minutes
17. Take the gel out and view the gel under a UV transilluminator for analysis.