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Contents |
Assessment of previous results
We took another look at the gel from Friday 3 August. The backbones look OK, but the PCR bands are of the wrong size. There has probably been no amplification. We ran a Nanodrop on the purified gel bands (QIAGEN Gel Extraction Kit).
Protocol: DNA Concentration Measurement
- Take a 6 µl aliquote of the DNA and put back the main DNA tube in the fridge.
- Go to the room by the E.Coli lab (LBTM, not on Friday morning!) with:
- The 6 µl aliquote
- A 10 µl pipet
- Optionally, the buffer you used for DNA elution (there might be some next to the machine).
- The machine is the NanoDrop Spectrophotometer.
- On the computer, click on "Nucleic Acid".
- Put a 2 µl drop of (nuclease-free) water on the machine's tip as you are asked to and measure.
- Clean tips (both sides) with a quarter of tissue.
- Add 2 µl of the buffer you use and click on "Blank".
- Clean tips (both sides).
- Add 2 µl of your DNA sample and click "Measure".
- Clean tips (both sides) with a tissue.
- Take 2 measurements per sample (for averaging).
- Print the report when you are done
- Click on exit.
The important numbers are:
- 260/280 ratio, must be > 1.8
- 260/230 ratio, must be > 2 (too big, > 2.5? , might mean too much salts)
- Of course the DNA concentration.
Nanodrop results :
| Sample | 260/280 | 260/230 | conc. ng/µL |
| 1. L4 (pMP w/NotI & SpeI) | 1.83 | 0.09 | 26.76 |
| 2. L5 (pGL4.30 w/HindIII & MfeI) | 1.92 | 0.03 | 14.98 |
| 3. L6 (pGL4.30 w/HindIII & FseI) | 1.88 | 0.05 | 26.38 |
| 4. GFP PCR product | 2.76, 1.69 | 0.01, 0.02 | 5.42, 8.91 |
| 5. TNFR PCR product | 4.49, 2.32 | 0.01, 0.01 | 2.58, 3.46 |
| 6. SEAP PCR product | 11.27, 1.86 | 0.01, 0.01 | 2.64, 6.23 |
New PCR of the readouts
Protocol: PCR
PCR is a reaction that makes it possible (and relatively easy) to amplify
a certain region of DNA. The first step is the selection of that region
(and the design of the relevant primers). Primer design can be done by hand, or by
using our Primer Design Helper. Once
done, order the primers (in our case, we ordered from them [http://www.idtdna.com/ IDT]).
When you've received the primers, prepare them and make sure you've got your PCR kit (we used the "Phusion® High-Fidelity DNA Polymerase"). Start preparing your master mix, the composition for one tube is:
1X Mastermix 20μl reaction, add in this order
| Reagent | Volume [μl] |
|---|---|
| Water | Complete to total volume of 20μl |
| HF-Buffer (5x) | 4 |
| DMSO (optional) | 0.6 |
| dNTPs | 0.4 |
| Forward primer (50μM) | 0.2 |
| Reverse primer (50μM) | 0.2 |
| Template (10ng/μl) | 0.5 |
| Phusion HF polymerase | 0.2 |
Prepare one or two extra tubes-worth of reagent (you'll use some liquid on the walls of your tips).
Once you've finished, you should run the resulting products on a gel to check if everything went as planned.
Tips
- Thaw the HF-Buffer, DMSO and dNTPs before making the mastermix.
- Avoid taking the Phusion-HF polymerase out of the freezer (only take it out briefly when you need to add it).
- If the reactions have different primers and/or template, add the polymerase right after the dNTPs, split the mastermix and add the rest.
- Don't forget positive and negative controls
- Primers should have similar Tms (less than 5°C).
- Primer Tm calculation is a less exact science than it should be (just test several tools and compare their results). If you're not sure what the correct Tm is, consider using a gradient PCR.
- Avoid primers with strong secondary structures.
- PCR can introduce mutations. Don't forget to sequence your final product (this could be your final plasmid): you really don't want to lose a few weeks because of a "corrupt" plasmid.
First, a primer dilution was made. Stock primers at 500 µM were brought to a 10 µM concentration. (1 µl primer + 49 µl H20)
Then, the PCR was re-run with a slightly adapted protocol (different temperatures and cycle durations, slightly different mix composition - Master Mix 2, rediluted primers). We also retried a PCR with the same primer with the master mix from the 3rd of August (called Master Mix 1 further on).
- Master Mix 2, for 4 tubes (3 + an excess one)
- H20: 110 µl
- Buffer HF (5x): 40 µl
- dNTP (10 mM): 4 µl
- Phusion DNA Polymerase: 2 µl
Then split into 3 tubes, add the following reagents to every one of them:
- Primers:
- Forward (50 µM): 5 µl
- Reverse (50 µM): 5 µl
- Template DNA: 1 µl
The total volume in each tube should be 50 µl.
- Thermal cycling protocol
- Initial denaturation: T = 98°C, 5 min
- Cycle through this 30 times:
- Denaturation: T = 98°C, 15 sec
- Annealing: T = 65°C, 30 sec
- Extension: T = 72°C, 1 min
- Final extension: T = 72°C, 10 min
- Hold: T = 4°C, stay until pickup
Nanodrop
Protocol: DNA Concentration Measurement
- Take a 6 µl aliquote of the DNA and put back the main DNA tube in the fridge.
- Go to the room by the E.Coli lab (LBTM, not on Friday morning!) with:
- The 6 µl aliquote
- A 10 µl pipet
- Optionally, the buffer you used for DNA elution (there might be some next to the machine).
- The machine is the NanoDrop Spectrophotometer.
- On the computer, click on "Nucleic Acid".
- Put a 2 µl drop of (nuclease-free) water on the machine's tip as you are asked to and measure.
- Clean tips (both sides) with a quarter of tissue.
- Add 2 µl of the buffer you use and click on "Blank".
- Clean tips (both sides).
- Add 2 µl of your DNA sample and click "Measure".
- Clean tips (both sides) with a tissue.
- Take 2 measurements per sample (for averaging).
- Print the report when you are done
- Click on exit.
The important numbers are:
- 260/280 ratio, must be > 1.8
- 260/230 ratio, must be > 2 (too big, > 2.5? , might mean too much salts)
- Of course the DNA concentration.
The DNA concentration after the PCR was found to be quite high, usually above 300 ng/µl.
Gel electrophoresis to check the size of the PCR products
Protocol: Gel Electrophoresis
Agarose concentration depends on the size of the DNA to be run. We will mostly use 1%.
VOL is the desired volume of gel in ml:
CH Lab
- Add 0.01*VOL g of agarose to a clean glass bottle.
- Pour VOL/50 ml of 50xTAE in a graduated cylinder. Fill up to VOL ml with di water.
- Add the resulting VOL ml of 1xTAE to the glass bottle with agarose.
- Microwave, at 7, the bottle (loose cap!) until it boils.
- Carefully remove bottle (can be super heated!) and check for the total absence of particles. Microwave again if needed.
- Prepare a gel box, with comb, and fill it up with the agarose solution (maybe not the whole solution is needed).
- Add 0.05 µl per ml of gel in the box of Red Gel (it's in the iGEM drawer) and stirr until disolved.
- Wait until cold and solidified.
- Carefully remove comb.
- Place the box in the electrophoresis chamber.
- Fill up the electrophresis chamber with 1x TAE buffer.
- Add blue dye to the DNA samples (6x loading buffer, that is 10 µl in 50 µl of DNA solution).
- Inject 30 µl of ladder marker in the first well (that's 1 µg of DNA).
- Inject 60 µl of each DNA solution in the other wells.
- Set voltage to 70-90 V and run for 30-40 min, or until the dye reaches the last 25% of the gel length (DNA travels from - to +).
- Place the gel under the camera, cover, turn UV on and take photos!
Preparing the ladder:
- get 1kb ladder DNA from the freezer (500 µg/ml).
- for 30 charges, 30 µl per charge, we need 900 µl:
- 60 µl of 1kb ladder DNA
- 150 µl of dye (6x loading buffer)
- 690 µl of water
BM Lab
In this lab the gels are slightly different. The total volumes for the small, the medium and the large gel are respectively 60ml, 80ml and 90ml. As we use 0.5x TAE buffer instead of 1x, we can use higher voltages (170V seems to work fine). The gel should run 20-40 minutes, not more. As the gel is thinner, load less DNA (up to ~10ul).
Made a 150 ml 1% gel with 12 wells, loaded in the following order:
- Lane 1: Ladder
- Lane 2: Master Mix 1
- Lane 3: TNFR 1 (1 stands for "with Master Mix 1")
- Lane 4: GFP 1
- Lane 5: SEAP 1
- Lane 6: Master Mix 2
- Lane 7: TNFR 2
- Lane 8: GFP 2
- Lane 9: SEAP 2
- Gel picture
We seem to have at least one working PCR (with a band) for each readout. This isn't a very high-quality PCR, though.
Digestion of the PCR products
Protocol: Restriction site digestion
- Look for the best pair of restriction sites, ideally with similar digestion temperatures and times.
- [http://tools.neb.com/NEBcutter2/ NEBcutter] for finding cutting enzymes.
- [http://www.neb.com/nebecomm/DoubleDigestCalculator.asp Double Digest Finder] for the parameters.
- Calculate the amounts required of:
- DNA
- Buffer (usually from 10x to 1x)
- BSA, if needed (usually from 100x to 1x)
- Enzymes (depends on the amount of DNA)
- Water
- Get the recommended buffer (and BSA if needed) from the freezer and let defreeze.
- Mix all the ingredients, except DNA, in a tube.
- Note: Enzymes should stay no longer than a couple of minutes out of the freezer. Don't touch the bottom of the tubes! Don't vortex!
- Distribute the mix in as many tubes as DNA samples and add the DNA.
- Keep in the Thermomixer at the recommended temperature.
Sowmya's recommended amounts (50 µl total solution):
- 5 µl of 10x buffer
- 0.5 µl of 100x BSA
- 1 µl of each enzyme
- 5 µl of DNA
- 37.5 (up to 50 µl) of water.
Protocol based on what was done on July the 4th.
The PCR products that were found acceptable were digested as follows:
- Digestion mixes
- GFP with FseI & HindIII:
- GFP PCR product: 40 µl
- FseI: 2 µl
- HindIII: 1 µl
- 10x BSA: 5 µl
- 10x N2 buffer: 5 µl
- H2O: 37 µl
- TNFR with FseI & HindIII:
- TNFR PCR product: 40 µl
- FseI: 2 µl
- HindIII: 1 µl
- 10x BSA: 5 µl
- 10x N2 buffer: 5 µl
- H2O: 37 µl
- SEAP with MfeI & HindIII:
- SEAP PCR product: 40 µl
- MfeI: 1 µl
- HindIII: 1 µl
- 10x N2 buffer: 5 µl
- H2O: 43 µl
The total digestion volume is slightly bigger than usual.
Ligation of the PCR products
Protocol: Ligation
Ligation is a method of combining several DNA fragments into a single plasmid. This is often the
step following a PCR (and a PCR cleanup) or a gel extraction. You can also do a "dirty" ligation, where you follow a certain number of digestions directly by a ligation.
- Download the following spreadsheet : File:Team-EPF-Lausanne Ligation.xls
- Fill in the pink areas with the vector and fragment concentration, their size and the ratio.
- Add all the suggested ingredients order in a microcentrifuge tube, in the order they appear.
- Ligate for 2 hours at 14ºC.
- Immediately transform competent bacteria with the ligation product.
Note: This protocol hasn't been optimized for blunt-end ligation (though it might still work).
We used three different fragment-to-vector ratios (1:2, 1:3, 1:4 and a 1:3 control without a fragment), because we were not sure of the Nanodrop concentrations. Eleven tubes were made:
- Tube 1: GFP + pGL4.30 (HindIII/FseI) 1:2
- Tube 2: GFP + pGL4.30 (HindIII/FseI) 1:3
- Tube 3: GFP + pGL4.30 (HindIII/FseI) 1:4
- Tube 4: nothing + pGL4.30 (HindIII/FseI) 1:3 CONTROL
- Tube 5: TNFR + pGL4.30 (HindIII/FseI) 1:2
- Tube 6: TNFR + pGL4.30 (HindIII/FseI) 1:3
- Tube 7: TNFR + pGL4.30 (HindIII/FseI) 1:4
- Tube 8: SEAP + pGL4.30 (HindIII/MfeI) 1:2
- Tube 9: SEAP + pGL4.30 (HindIII/MfeI) 1:3
- Tube 10: SEAP + pGL4.30 (HindIII/MfeI) 1:4
- Tube 11: nothing + pGL4.30 (HindIII/MfeI) 1:3 CONTROL
The ligation tubes were than incubated for two hours at 14°C.
Transformation
Protocol: E.Coli Transformation
- Thaw the competent E.coli (DH5alpha) cells on ice (not in hands!)
- As soon as it is thawed, add 50µl of the cells to the DNA (~50-100 ng of pure plasmid, or some 2 µl usually)
- Let it rest on ice for 20-30 min. Meanwhile, put agar plate (with correct antibiotic) at 37°C for prewarming.
- Put the tube with DNA+E.coli at 42°C for 45 sec - 1 min (heat shock)
- Add 400 µl of LB broth and place at 37°C for 20-30 min (shaking)
- Spread the cells on the prewarmed plate (and let it dry)
- Incubate the plate upside-down at 37°C for ~14-15 hours (leaving it more than 16h decreases the plasmid quality)
Bacteria were transformed with the ligation mixes, and then plated (11 plates).
