Difference between revisions of "Team:elan vital korea/Protocol"

We conducted our experiments by following the protocols below. As an official procedure, lab workers should understand the lab experiment assigned to them along with safety procedures before starting lab work. The protocols are arranged according to the order of experiments we followed.

1. Reporter cell, test cell and competent cell (Top 10 invitrogen) must be kept at 4°C and frequently used reagents, reagents, DNA plasmids should be kept at −20°C in the freezer.

2. We use triple distilled water (or DDH2O) to make LB broth. Triple distilled water is kept at lab temperature (around 18 °C or lower).

3. Other materials such as yeast and NaCl are stored and maintained under the responsibility of Gachon Molecular Biology Lab.

4. We have to record the history of each material, including if plasmids/reporter cell/ test cell/ AHL have been frozen and if so, when it is used.

1. To prevent contamination, we only used LB medium made within three days.

2. Materials: Sodium Chloride (LB Media, Sigma), Trypton(LB Media, Sigma), Yeast Extract(LB Media Sigma), ddH2O (triple distilled water)

3. Equipment: autoclave, electronic scale.

4. Protocol For 200mL LB bottle
1) 2 g of Sodium Chloride to a final concentration of 0.17 M
2) 2g of 1%(w/v) Bacto™ tryptone
3) 1g of 0.5% (w/v) yeast extract
4) ddH2O to 200 mL
5) Autoclave for 20 min within 2 hours
6) Keep at room temperature

1. We have used LB (solidified lysogeny broth), rich growth medium for E.coli, in our experiments.

2. Just before pouring the solution into petri dishes, an antibiotic can be added for resistance selection. We followed the normal working concentrations such as:
- chloramphenicol: 25 μg/mL (Chloramphenicol stock is dissolved in ethanol) In case of using ampicillin: 100 μg/mL
- normal stock concentrations:1000-fold

3. Material to make LB plates: Sodium Chloride (LB Media, Sigma) Bacto™ tryptone (LB Media, Sigma) yeast extract (LB Media, Sigma) Bacto™ agar (LB Media, Sigma) ddH2O (triple distilled water) 1000x chloramphenicol or ampicillin

4. LB agar preparation protocol We usually make 1liter bottle for LB Agar
1) 200 mL LB prepared fresh, non-autoclaved
2) 3 g agar
3) Shake until all solids are dissolved
4) Autoclave for 20 min within 2 hr
5) Keep it cool until it reaches around 40-50 °C
6) Add 200 μL of 1000x chloramphenicol and gently stir it. Be careful not to shake the bottle too long/hard so that bubbles are created.
7) Pour into empty petri dishes just enough to cover the surface (~20 mL per plate). In case that bubbles are in the plate, heat the plate surface carefully with a burner only until the bubbles are burst but the solution is heated.
8) Leave the plates at room temperature around one hour until it is solidified.
9) Solidified plates should be turned upside down for a few hours at room temperature, then stored at 4°C.

1. We have conducted overnight culture for a single bacterial strain which process needs a plate or medium with single colonies and LB containing chloramphenicol.

2. Material Needed chloramphenicol: 25 μg/mL Normal stock concentrations: 1000-fold higher In case of using ampicillin: 100 μg/mL

3. Protocol
1) Quickly burn the neck of a bottle containing LB medium before pouring it out into a tube. Even the slightest contamination of LB will be damaging.
2) Add chloramphenicol or ampicillin to give the appropriate concentration
3) Scoop one colony from the plate with a sterile micropipette tip
4) Immediately stick the tip into the tube containing the medium and chloramphenicol or ampicillin
5) Incubate at 37°C with the shaking incubator overnight.

1. Agarose gel electrophoresis is used for separation and analysis of larger (>100 bases in length) nucleic acids under non-denaturing conditions.

2. Analysis requires that the gel contains a DNA stain visible under UV light. Since the stain interacts with nucleic acids and is therefore potentially mutagenic, always wear nitrile gloves when working with agarose gels.

3. Use protective glasses when using the UV light box.

4. Material Needed
Agarose
1x TBE
Sybr®Safe
Loading dye mix
DNA ladder size marker
DNA samples

5. Protocol:
1) The gel tray must be on a level surface.
2) Insert the comb into the gel tray at one end ~1 cm from the edge.
3) For a 1% 150 mL agarose gel, weigh 1.5 g of agarose in a 500 mL conical flask.
4) Add 150 mL 1x TBE buffer.
5) To dissolve the agarose in the buffer, swirl to mix and microwave for a few minutes taking care not to boil the solution out of the flask.
6) Remove the flask occasionally and check whether the agarose has dissolved completely.
7) Let the agarose solution cool down.
8) Once the solution is touchable, add the DNA stain.
9) Check the stock concentration as the working concentration for ethidium bromide is 0.5 μg/mL while for Sybr®Safe it is simply 1x.
10) Pour the gel solution into the gel tray.
11) Remove any air bubbles with a pipette tip.
12) Put in comb.
13) The gel will solidify while cooling down to room temperature, which usually takes about 30 min.
14) Running the gel by the following procedure
a. Release the gel tray from the tape or casting stand.
b. Place the gel tray into the buffer chamber and remove the comb carefully
c. Add 1x TBE buffer until the gel is completely covered.
d. Take the DNA sample (~0.2 μg) and mix with loading dye.
e. Load the size marker mixed in 1x loading dye (~6 μL final volume) into a middle well.
f. Load the samples into the other wells while writing down which lanes have which samples.
g. Put the lid onto the buffer chamber and connect it to the power supply.
h. Run the gel at 100 V for 30–60 min. Neither of the two dyes should be run off the gel.
i. Stop the run and bring the gel to a UV table to visualize the gel bands.
j. Take a picture of the gel.

1. This protocol is for the purification of up to 10μg DNA (70bp to 10kb).

2. The yellow color of buffer QG indicates a pH ≤ 7.5. DNA adsorption to the membrane is only efficient at pH ≤ 7.5.

3. Add ethanol (96%100%) to Buffer PE before use (see bottle label for volume).

4. Isopropanol (100%) and a heating block or water bath at 50°C are required.

5. All centrifugation steps are carried out at 17,900 x g (13,000 rpm) in a convetional table-top microcentrifuge.

6. Symbosl: ● centrifuge processing; ▲ vacuum processing.
1. Excise the DNA fragment from the agarose gel with a clean, sharp scalpel.
2. Weigh the gel slice in a colorless tube. Add 3 volumes Buffer QG to 1 volume gel (100 mg gel ~ 100μl). The maximum amount of gel per spin column is 400mg. For >2% agarose gels, add 6 volumes Buffer QG.

3. Incubate at 50°C for 10 min (or until the gel slice has completely dissolved). Vortex the tube every tube every 2-3 min to help dissolve gel. 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 10μl 3 M sodium acetate, pH 5.0, and mix. The mixture turns yellow.
4. Add 1 volume isopropanol to the sample and mix.
5. Place a QIAquick spin column in ● a provided 2ml collection tube or into ▲ a vacuum amnifold. To bind DNA, aply the sample to the QIAquick column and ● centrifuge for 1 min or ▲ apply vaccum to the manifold untill all the samples QIAquick column back into the same tube. For example volumes of 0> 800μl, load and spin/apply vacuum again.
6. If the DNA will subsequently be used for sequenceing, in vitro transcruption, or microinjection, add 500μl Buffer QG to the QIAquick column and ● centrifuge for 1 min or ▲ apply vaccum. ● Discard flow through and place the QIAquick column back into the same tube.

7. To wash, add 750μl Bufick column and fer PE to QIAquickcolumn and ● centrifuge for 1 min or ▲ apply vacuum. ●Discard flow-through and place the QIAquick column back into the same tube.