IGEM:Harvard/2008/Lab Notebooks/GenProtocols

QIAprep Spin Miniprep Kit

This protocol is designed for the purification of up to 20 ug high-copy plasmid DNA from 1-5 ml overnight E. coli culture in LB medium.

Procedure

1. Resuspend pelleted bacterial cells in 250uL Buffer P1 and transfer to a microcentrifuge tube.
2. Add 250 uL Buffer P2 and mix thoroughly by inverting the tube 4-6 times.
3. Add 350 uL Buffer N3 and mix immediately and thoroughly by inverting the tube 4-6 times.
4. Centrifuge for 10 min at 13,000 rpm in a table-top microcentrifuge.
5. Apply the supernatant to the QIAprep spin column by pipetting
6. Centrifuge for 30-60 s. Discard flow-through.
7. Wash QIAprep spin column by adding 0.75 ml BUffer PE and centrifuging for 30-60s.
8. Discard the flow-through, and centrifuge for an additional 1 min to remove residual wash buffer.
9. To elute DNA, place the QIAprep column in a clean 1.5 ml microcentrifuge tube. Add 50 uL Buffer EB to the center of each QIAprep, let stand for 1 min, and centrifuge for 1 min.

Ligation protocol, using Roche Rapid Ligation Kit

Brief Protocol

Mix:

• 5x DNA dilution buffer – 2uL
• Vector (cut and CIP treated) – 0.25-1uL
• Insert DNA – 0.5-6.5 uL
• ddH2O – to final volume of 10uL

Vortex and quick centrifuge above mixture, then add:

• 10uL of 2X rapid ligation buffer
• 1 uL ligase

Vortex and quick centrifuge again. Hold at room temp ~20 minutes, then use 5uL of the ligation reaction to transform 50uL of chemically competent E. coli (eg, TOP10 or DH5-alpha).

Protocol Notes

Reagents

Make sure buffers are completely thawed before use. The 2x rapid ligation buffer contains a reducing agent that looks like white flakes when thawed. These white flakes must be completely dissolved back into the buffer before use. Keep the ligase enzyme at -20 degrees until just before you are ready to add it, and return it to the freezer promptly. Minimizing time outside of the freezer will preserve the activity of the enzyme.

DNA quantities to use

The ratio of insert DNA to vector is very important. Ideally, you want 10 units of insert to each 1 unit of vector. To estimate this, use the Nanodrop to quantitate the amount of vector and insert you have. Note, the Nanodrop will give you concentrations in nanograms, so you will need to roughly convert this to a ratio based on the size of the insert and the vector.

Example: Vector DNA is 4000 base pairs, and you have 100ng per uL concentration. You use 0.5 uL (50ng) of vector. Your insert is 800 base pairs, and you have 50ng per uL. Your vector is 5 times as large as your insert, which means that on a per-nanogram basis, there are 5 times as many individual units of insert per nanogram in your insert sample than in your vector sample. So, you should use only double the nanogram quantity of insert (2uL) in this reaction (5x more units per ng X 2uL = 100ng = 10x insert compared to vector). As you can see, this is a very rough calculation, but it works pretty well.

Troubleshooting

What to try if your ligation isn’t working (in this order). Foremost among these is to ask your TFs for help and advice. Ask for help sooner rather than later to avoid headaches!

1. Repeat the ligation. Set up three reactions, one using 0.5uL of insert, one with 2uL of insert and one with the maximal volume the reaction will allow (7-7.5uL insert). Do all of these reactions in parallel. Sometimes, the above mentioned rough quantitation scheme doesn’t work, so it is best to try a range of insert to vector ratios.
2. Review all of your digestion steps – are you sure you cut the vector and the insert with the correct enzymes? Did you let it digest for long enough? Did you CIP treat the vector after the digest by before the purification? (Many people don’t use CIP on their cut vectors, but I find it increases the success of your cloning endeavours enormously). If you have any doubts about the digestion steps, do them again and then repeat the ligation with three quantities of insert, as described above.
3. Rarely, enzymes do go bad (perhaps a labmate unintentionally left the enzyme at room temp for too long and then put it back). Try a new kit.
4. Still not working? Send the parts you are trying to clone for sequencing. Often, DNA obtained from other scientists or from the parts registry is contaminated or just plain wrong. You may not be cloning what you think you are cloning.

Bacterial Transformation Protocol

Chemically competent cells

To make chemically competent cells

Day 1:

1. Grow 5mL culture of bacteria in LB media overnight, 37° C , shaking

Day 2:

1. Inoculate fresh 100mL LB culture with 1mL bacteria from 5mL culture
2. Grow culture at 37°, shaking, to an OD600 of 0.2-0.4 (takes 1-3h.)
   Perform the remainder of the protocol in the cold room OR take care to make sure cells remain cold!!!
3. Decant culture into two 50mL falcon tubes and chill on ice 15 minutes.
4. Spin for 15 minutes at 2500 RPM, 4°C
5. Remove supernatant and wash pellets in 50 mL ice cold 100mM MgCl2 (add 25 mL per tube and combine tubes)
6. Spin for 15 minutes at 2500 RPM, 4°C
7. Remove supernatant and resuspend pellet in 50mL ice cold 100mM CaCl2
8. Incubate on ice 30 minutes
9. Spin for 15 minutes at 2500 RPM, 4°C
10. Remove supernatant and gently resuspend pellet in 10 mL ice cold 100mM CaCl2 + 15% glycerol (v/v).
11. Incubate on ice 30 minutes.
12. Aliquot 100uL to 0.6mL tubes and store at -80°C. (makes ~100 aliquots)

To transform chemically competent cells

1. Soak the spots in 5 µL of the warmed TE for 20 minutes. This allows the maximum concentration of DNA in solution. Start thawing the competent cells on wet crushed ice.
2. Chill labeled 2 ml conical bottom tubes on wet ice. Add 2 µL of DNA in TE and 50 µL of thawed TOP10 competent cells to the tubes. In our experience, these volumes have the best transformation efficiency. The 2 ml tubes allow better liquid movement during incubation. Extra eluted DNA may be held at least several weeks frozen or at refrigerator temperature.
3. Hold the DNA and competent cells on ice for 30 minutes. This improves transformation efficiency by a significant amount.
4. Heat shock the cells by immersion in a pre-heated water bath at 42ºC for 60 seconds. A water bath is important to improve heat transfer to the cells.
5. Incubate the cells on ice for 2 minutes.
6. Add 200 μl of SOC broth (check that this broth is not turbid, which would indicate previous contamination and bacterial growth). This broth should contain no antibiotics.
7. Incubate the cells at 37ºC for 2 hours while the tubes are rotating or shaking. We have found that growth for 2 hours helps in transformation efficiency, especially for plasmids with antibiotic resistance other than ampicillin.
8. Label an LB agar plate containing the appropriate antibiotic(s) with the part number, plasmid, and antibiotic resistance. Plate 250 µl of the incubated cell culture on the plate.
9. Incubate the plate at 37ºC for 12-14 hours, making sure the agar side of the plate is up. If incubated for too long the antibiotics, especially ampicillin, start to break down and un-transformed cells will begin to grow.

For electrocompetent cells

To make electrocompetent cells

1. Grow bacteria to log phase (OD 0.4-0.6)
2. Aliquot 1mL of culture to a 1.5mL epi tube. Make as many aliquots as you need for the number of plasmids you need to transform
3. Centrifuge aliquots at 12,000g for 1 minute
4. Remove media and wash with 0.33 volumes (330μL) of 1M sorbitol
5. Centrifuge at 12,000g for 1 minute
6. Remove wash and resuspend in 0.05 volumes (40μL) 1M sorbitol.
7. Place tubes in ice and use within 15 minutes.
8. Add 100-500ng of plasmid DNA to the tube
9. Electroporate at 0.55kV