Difference between revisions of "Team:KU Leuven/Research/Methods"

Theory
To be able to create patterns, two different cell types called A and B will interact with each other. In order to achieve the desired behavior, the cells used in the experiments were derived from K12 Escherichia coli strains with introduction of specific knockouts. Cell type A has a deletion of tar and tsr, whereas in cell type B both tar and cheZ are knocked out. The Keio collection is composed of a set of precisely defined single-gene deletions of all nonessential genes in E. coli K-12. The targeted genes were replaced by a kanamycin resistance cassette. The kanamycin cassette is enclosed between two FRT sites making excision possible using FLP recombinase (reference 1). FLP recombinase triggers an intramolecular recombination between FRT repeats in the chromosome. Since both the antibiotic resistance gene and the plasmid replication region are surrounded by two FRT sites both are to be eliminated (Figure 1, step 1).

A genetic procedure for moving selectable mutations of interest called the P1 transduction was used. Since the packaging of the bacteriophage P1 is rather inaccurate, it will on occasion package the DNA of its bacterial host instead of its own phage chromosome. This implies that the lysate contains either packaged phage or bacterial DNA. After infection of a second host with this lysate, a transfer of parts of the chromosome from the donor strain into the receiver strain will take place. Those DNA pieces can then recombine using the FRT sites and hereby be incorporated permanently into the chromosome of the new strain. Here, the recombination was triggered by selection on kanamycin. (reference 2)

In general, we used three steps to obtain our double knock-outs (Figure 1). In the first step, the kanamycin cassette of our tar knock-out strain was removed by flippase, coded on plasmid PCP20. Afterwards, the temperature sensitive plasmid was removed by growing the cells overnight at 42°C. In a third step, the tar knock-out strain is infected by lysate originating from our tsr and cheZ knock-out strains. After selection on kanamycin plates, we obtained our double knock-outs. These knock-outs were confirmed by PCR. For more information, please check our result page.

Figure 1

P1 transduction. Click to enlarge

Protocol

1. Preparation of lysate starting from stock plate of phage

1. Make an overnight culture of E. coli MG1655.

2. Take 500 µL overnight culture and add the phage P1. Incubate overnight at 37°C.

3. Take single plaques of the P1 stock plate and bring this in a sterile Eppendorf tube together with 200 µL of mQ.

4. Overnight extraction while shaking at 37°C.

5. Add 0.01, 0.1, 10 and 100 µL of extraction to 500 µL of a stationary phase culture of E. coli MG1655. Vortex and plate out.

6. Add LB soft agar containing 10 mM MgSO₄ and 5 mM CaCl₂ and incubate at 37°C.

7. Choose the plate with the best lysates.

8. Sterilize your spoon using a Bunsen burner, cool it down with water and wash it with 100% ethanol.

9. Cut out a plaque from the soft agar and put this in a 10 mL syringe.

10. Press the content of the syringe in an Eppendorf tube and centrifuge for 10 minutes at 14000 rpm.

11. Take 650 µl and bring this in a new Eppendorf tube.

12. Extraction with 30 µL of CHCl₃.

13.Vortex vigorously.

14. Store lysate at 4°C.

2. Preparation of the lysate of donor strain

1. Firstly, centrifuge the lysate to ensure the chloroform is at the bottom of the Eppendorf tube. Then add 0.1, 1, 10 and 100 µl of lysate to 500 µL stationary phase overnight culture of the donor strain.

2. Add LB soft agar containing 10 mM MgSO₄ and 5 mM CaCl₂. Incubate this at 37°C.

3. Sterilize your spoon in a Bunsen flame, cool it down with water and wash with 100% ethanol.

4. Centrifuge the Eppendorf tubes 10 minutes at 14000 rpm.

5. Transfer 650 µL into a new Eppendorf tube.

6. Extract with 30 µL of CHCl₃.

7. Vortex vigorously.

8. Store the lysate at 4°C.

3. Transduction to acceptor strain

1. Concentrate 500 µL of stationary phase overnight acceptor strain culture five times in LB with 10 mM MgSO₄ and 5 mM CaCl₂.

2. Add 0.1, 1, 10 and 100 µL of donor strain lysate to 100 µL acceptor strain.

3. Incubate 30 minutes at 37°C.

4. Plate out on selective medium and incubate overnight.

5. Plate out lysate-only to check for contamination as well.

Theory
The Gibson assembly as described by Gibson et al., is a rapid DNA assembly method which assures directionional cloning of fragments in one single reaction. For the Gibson assembly to happen, three essential enzymes are needed: a mesophylic nuclease, a thermophylic ligase and a high fidelity polymerase. For this reaction, we used NEBuilder. In the first step of this reaction, the exonuclease rapidly cleaves off the 5’ DNA ends. The exonuclease is unstable at 50°C and gets degraded. In the second step, the designed sequence overlaps anneal and the polymerase starts filling in the gaps. In the final step, the ligases covalently joins both ends. After this, the plasmid is ready to be transformed. This text was based on the IDT website as seen on 13/09/2015

Figure 1

Gibson assembly reaction and its essential components E.coli

Materials

Protocols

Protocol

1. Prepare selective media (LB with 0.25% agar (2,5 g/L) in Petri dishes (85 mm dia).

2. Apply 1.5 µL of the diluted cell suspensions from mid-log-phase cultures (~2×105 cells/µL (OD=0.5)) to the center of the plates, and let dry in air for 15 min.

3. Incubate at 37 °C for 10 h.

Theory

Tekst

Protocol

Make a standard curve

The goal is to measure the absorbance of violacein versus the concentration of AHL around 0.04 nm.

1. Prepare selective media (LB with 0.25% agar (2,5 g/L) in Petri dishes (85 mm dia).

2. Apply 1.5 µL of the diluted cell suspensions from mid-log-phase cultures (~2×105 cells/µL (OD=0.5)) to the center of the plates, and let dry in air for 15 min.

3. Incubate at 37 °C for 10 h.

Preparation of media

The goal is to measure the absorbance of violacein versus the concentration of AHL around 0.04 nm.

1. Prepare selective media (LB with 0.25% agar (2,5 g/L) in Petri dishes (85 mm dia).

2. Apply 1.5 µL of the diluted cell suspensions from mid-log-phase cultures (~2×105 cells/µL (OD=0.5)) to the center of the plates, and let dry in air for 15 min.

3. Incubate at 37 °C for 10 h.

Quantification

The goal is to measure the absorbance of violacein versus the concentration of AHL around 0.04 nm.

1. Prepare selective media (LB with 0.25% agar (2,5 g/L) in Petri dishes (85 mm dia).

2. Apply 1.5 µL of the diluted cell suspensions from mid-log-phase cultures (~2×105 cells/µL (OD=0.5)) to the center of the plates, and let dry in air for 15 min.

3. Incubate at 37 °C for 10 h.

Materials

- 1L sterile LB without NaCl (10g tryptone, 5g yeast extract per 1L)

- 500 mL of 10% v/v glycerol

- Cold falcon tubes of 50 mL

- Cold eppies and pipette tips

Protocol

Day 1:

1. Strike the cells on a plate and grow overnight at 37°C.

Day 2

1. Pick a single colony from your plate and grow it in 1-3 mL salt free LB overnight at 37°C.

Day 3:

1. Grow 300-400 mL cells (without salt) in 37°C until the OD reaches 0.6 (use a starting culture).

2. Cool down on ice and from now on perform all the steps at 4°C.

3. Spin the cells down in falcon tubes (3500 g, 20 min, 4°C). Using falcon tubes ensures no detergents present.

4. Resuspend the pellet in 30 mL icecold 10% glycerol (filtered to a disposable bottle to ensure the absence of detergents). Spin down the cells (5000 g, 10 min, 4°C). Repeat this step 3 times.

5. Resuspend the cells in 10% glycerol to obtain a dense pulp (usually not more than 1.5 mL).

6. Take 50 µL sample and do the electroporation test (without DNA). You should have a pulse of 4-6 msec. If shorter, wash the cells once again with 30 mL glycerol.

7. Aliquot the cells (50 µL), quick-freeze in liquid nitrogen and store at -80°C.

Materials:

- DNA

- Electrocompetent cells

- SOC medium

- Ice-cold cuvettes

Protocols:

1. Add 1 µL DNA to 50 µL of electrocompetent cells in an ice-cold cuvette (1 mm).

2. Electroporate (Eppendorf, 1700 V, 4 msec).

3. Add 950 µL of SOC solution.

4. Incubate for one hour at 37°C.

5. Plate 50 µL out on pre-warmed plates (37°C).

Materials:

Overnight liquid cultures

Thermo Scientific geneJET Plasmid Miniprep Kit

Protocol :

1. Transfer the liquid cultures to an Eppendorf tube and spin down at 13400 rpm for 1 to 2 minutes

2. Resuspend the pelleted cells in 250 µL of the Resuspension solution. The cells should be resuspended completely by vortexing or pipetting up and down until no cell clumps remain.

3. Add 250 µL of the Lysis Solution and mix thoroughly by inverting the tube 4-6 times until the solution becomes viscous and slightly clear.
Note. Do not vortex to avoid shearing of chromosomal DNA. Do not incubate for more than 5 min to avoid denaturation of supercoiled plasmid DNA.

4. Add 350 µL of the Neutralization Solution and mix immediately and thoroughly by inverting the tube 4-6 times.
Note. It is important to mix thoroughly and gently after the addition of the Neutralization Solution to avoid localized precipitation of bacterial cell debris. The neutralized bacterial lysate shoud become cloudy.

5. Centrifuge for 5 min to pellet cell debris and chromosomal DNA.

6. Transfer the supernatant to a supplied GeneJET spin column by decanting or pipetting. Avoid disturbing or transferring the white precipitate.
Note. Close the bag with GeneJET Spin Columns tightly after each use!

7. Centrifuge for 1 min. Discard the flow-through and place the column back into the same collection tube.

8. Add 500 µL of the Wash Solution (diluted with ethanol prior to first use) to the GeneJET spin column. Centrifuge for 30-60 seconds and discard the flow-through. Place the column back into the same collection tube.

9. Repeat the washing procedure (step 8) using 500 µL of the Wash Solution.

10. Discard the flow-through and centrifuge for an additional minute to remove residual Wash Solution. This step is essential to avoid residual ethanol in plasmid preps.

11. Transfer the GeneJET spin column to a fresh 1.5 mL microcentrifuge tube. Add 50 µL of the Elution Buffer to the center of the GeneJET spin column membrane to elute the plasmid DNA. Take care not to touch the membrane with the pipette tip. Incubate for 2 min at room temperature and centrifuge for 2 min.
Note. An additional elution step (optional) with Elution Buffer or water will recover residual DNA from the membrane and increase the overall yield by 10-20%. For elution of plasmids or cosmids >20 kb, prewarm Elution Buffer to 70°C before applying to silica membrane.

12. Discard the column and store the purified plasmid DNA at -20°C.

Materials:

PCR mixture

Thermo Scientific GeneJET PCR Purification Kit

Protocol:

1. Add a 1:1 volume of Binding Buffer to completed PCR mixture and 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 µL of 3 M sodium acetate, pH 5.2 solution and mix. The color of the mix will become yellow.

2. Transfer up to 800 µL of the solution from step 1 to the GeneJET purification column. Centrifuge for 30-60 s. Discard the flow-through.
Note. If the total volume exceeds 800 µL, the solution can be added to the column in stages. After the addition of 800 µL 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. Close the bag with GeneJET Purification Columns tightly after each use!

3. Add 700 µL of Wash Buffer to the GeneJET purification column. Centrifuge for 30-60 s. Discard the flow-through and place the purification column back into the collection tube.

4. Transfer the GeneJET purification column to a clean 1.5 mL microcentrifuge tube, add 50 µL of Elution Buffer to the center of the GeneJET purification column membrane and centrifuge for 1 min.
Note. For low DNA amounts, the elution volume can be reduced to increase DNA concentration. An elution volume between 20-50 µL does not significantly reduce the DNA yield. However, elution volumes less than 10 µL are not recommended. If DNA fragment is >10 kb, prewarm Elution Buffer to 65°C before applying to column. If the elution volume is 10 µL and DNA amount is ≥5 µg, incubate column for 1 min at room temperature before centrifugation.

5. Discard the GeneJET purification column and store the purified DNA at -20°C.

Protocol :

1. Excise the DNA fragment from the agarose gel with a clean, sharp scalpel. Minimize the size of the gel slice by removing extra agarose.

2. Weigh the gel slice in a colorless tube. Add 3 volumes of Buffer QG to 1 volume of gel (100 mg ~ 100 µl).
Note. For >2% agarose gels, add 6 volumes of Buffer QG. The maximum amount of gel slice per QIAquick column is 400 mg; for gel slices >400 mg use more than one QIAquick column.

3. Incubate at 50°C for 10 min (or until the gel slice has completely dissolved). To help dissolve gel, mix by vortexing the tube every 2–3 min during the incubation. IMPORTANT: Solubilize agarose completely. For >2% gels, increase incubation time. After the gel slice has dissolved completely, check that the color of the mixture is yellow (similar to Buffer QG without dissolved agarose). If the color of the mixture is orange or violet, add 10 µl of 3 M sodium acetate, pH 5.0, and mix. The color of the mixture will turn to yellow. The adsorption of DNA to the QIAquick membrane is efficient only at pH ≤7.5. Buffer QG contains a pH indicator which is yellow at pH ≤7.5 and orange or violet at higher pH, allowing easy determination of the optimal pH for DNA binding.

4. Place a QIAquick spin column in a provided 2 ml collection tube. To bind DNA, apply the sample to the QIAquick column, and centrifuge for 1 min. The maximum volume of the column reservoir is 800 µL. For sample volumes of more than 800 µL, simply load and spin again.

5. Discard flow-through and place QIAquick column back in the same collection tube. Collection tubes are re-used to reduce plastic waste.

6. To wash, add 0.75 mL of Buffer PE to QIAquick column and centrifuge for 1 min. Note: If the DNA will be used for salt sensitive applications, such as blunt-end ligation and direct sequencing, let the column stand 2–5 min after addition of Buffer PE, before centrifuging.

7. Discard the flow-through and centrifuge the QIAquick column for an additional minute at ≥10,000 x g (~13,000 rpm). IMPORTANT: Residual ethanol from Buffer PE will not be completely removed unless the flow-through is discarded before this additional centrifugation.

8. Place QIAquick column into a clean 1.5 mL microcentrifuge tube.

9. To elute DNA, add 50 µL of Buffer EB (10 mM Tris·Cl, pH 8.5) to the center of the QIAquick membrane and centrifuge the column for 1 min at maximum speed. Alternatively, for increased DNA concentration, add 30 µl elution buffer to the center of the QIAquick membrane, let the column stand for 1 min, and then centrifuge for 1 min. IMPORTANT: Ensure that the elution buffer is dispensed directly onto the QIAquick membrane for complete elution of bound DNA. The average eluate volume is 48 µl from 50 µl elution buffer volume, and 28 µl from 30 µl. Elution efficiency is dependent on pH. The maximum elution efficiency is achieved between pH 7.0 and 8.5. When using water, make sure that the pH value is within this range, and store DNA at –20°C as DNA may degrade in the absence of a buffering agent. The purified DNA can also be eluted in TE (10 mM Tris·Cl, 1 mM EDTA, pH 8.0), but the EDTA may inhibit subsequent enzymatic reactions.

Protocol:

1. Measure Blank using the nanometer with 2 µL the buffer you used to elute your sample.

2. Put 2 µL of the sample on the nanometer, and average the result.