1. Combine stocks and adjust pH to 7.5 (use 1.0N HCl).
2. Aliquot into flasks (50 ml/125 ml flask) with cotton stoppers on top and autoclave.
3. After autoclaving and cooling, the pH may change, so it must be monitored.
4. For solid media, add 1% noble agar.
5. For BG-11 don't add NaNO3.
TRANSFORMATIONS OF CYANOBACTERIA (OBTAINED FROM STONY BROOK IGEM TEAM)
1. Measure the OD750 (must be approx. 0.7)
2. 15mL per transformation must be centrifuged at maximum speed for 10 min at room temperature.
3. Remove the supernatant by pipetting.
4. Resuspend pellet in 10 mL of 10 mM NaCl solution and centrifuge at maximum speed for 10 minutes.
5. Resuspend the pellet in 0.3 mL of BG-11 and transfer to a microcentrifuge tube.
6. Add between 50ng and 2ug of DNA.
7. Wrap the microcentrifuge tube in aluminum foil to protect from light and incubate at 30°C for 24 hours.
8. Plate the entire volume (0.3mL) of cells on the big plates containing antibiotics by streaking.
TRANSFORMATIONS OF CYANOBACTERIA (INVITROGEN)
1. Measure the optical density of the Synechococcus elongatus cultures (from step 10, page 13) at 750 nm (i.e., OD750).
Note: For best performance, the OD750 of cultures should be greater than 1 and less than 2.
2. Harvest 1.5 mL of the cells (per transformation) by centrifugation at 14,000 rpm for 3 minutes at room temperature.
3. Remove the supernatant by pipetting.
4. Resuspend the cells in 1 mL of GibcoTM BG-11 medium by gently pipetting up and down.
5. Centrifuge the cells at 14,000 rpm for 1 minute at room temperature, and remove the supernatant by pipetting.
6. Resuspend the cells in 100 μL of GibcoTM BG-11 medium by gently pipetting up and down.
7. Add 100 ng of supercoiled plasmid DNA (i.e., a pSyn_6 construct containing your gene of interest) into the resuspended cells. In a separate tube, add 100 ng of an empty pSyn_6 vector as a negative control.
8. Mix the DNA-cell suspension gently by flicking the tube.
9. Incubate the cell-DNA mixtures in the 34°C water bath with a dark lid for 4 hours. After the incubation is complete, remove the tubes from the water bath and wipe them with 70% ethanol.
10. Plate 80 μL and 5 μL of each transformation mixture on separate BG-11 agar plates containing 10 μg/mL of spectinomycin and pre-warmed to room temperature.
11. Place the plates with agar side down on illuminated shelves at room temperature (25–30°C). Do not stack the plates to ensure continuous and even illumination.
12. Incubate the plates for 5–7 days or until the colonies are ready to pick. The results from the transformation with the pSyn_6 construct will depend on the nature of your gene of interest.
LB BROTH PREPARATION
1. Add 37g of nutrient agar to 400 mL of the distilled water
PREPARATION OF LURIA BERTANI BROTH, MILLER
1. Suspend 25 grams of the Luria Bertani broth in 1000 ml distilled water
2. Heat if necessary to dissolve the medium completely
3. Sterilize by autoclaving at 15 lbs pressure (121°C) for 15 minutes
4. Dispense as desired
PREPARING CHEMICALLY COMPETENT E.COLI
1. Prepare 0.1M CaCl2 and 0.1M CaCl2 + 15% glycerol solutions beforehand
2. Take 1 colony of DH5-alpha strain of E.coli from LB plates and inoculate in 10mL of LB in 50 ml Falcon tube
3. Place the tube with DH5-alpha E.coli strains in LB into the shaking incubator at 250 rpm at 37℃, until OD600 reaches 0.2-0.3.
4. Once the OD600 reaches 0.2-0.3, place the tube with DH5-alpha strain on ice for 15 min. Keep in ice solutions of 0.1M CaCl2 and 0.1M CaCl2 + 15% glycerol, too.
5. Centrifuge cells at 4oC for 10 min.
6. Resuspend pellet with 3 ml of 0.1M CaCl2 and put on ice for 30 min.
7. Centrifuge cells again and resuspend in 300 µL of 0.1M CaCl2 + 15% glycerol.
8. Prepare 50 µL aliquots of resulting solution in separate Eppendorf tubes.
9. Place aliquots in a Cold room at -80℃.
was carried out according to IGEM Protocols
1. Resuspend DNA in selected wells in the Distribution Kit with 10µl dH20. Pipet up and down several times, let sit for a few minutes. Resuspension will be red due to the cresol red dye.
2. Label 1.5ml tubes with a part name or well location. Fill lab ice bucket with ice, and pre-chill 1.5ml tubes (one tube for each transformation, including your control) in a floating foam tube rack.
3. Pipette 50µl of competent cells into the 1.5ml tube: 50µl in a 1.5ml tube per transformation. Tubes should be labeled, pre-chilled, and be in a floating tube rack for support. Keep all tubes on ice. Don’t forget a 1.5ml tube for your control.
4. Pipette 1µl of resuspended DNA into the 1.5ml tube: Pipette from well into the appropriately labeled tube. Gently pipette up and down a few times. Keep all tubes on ice.
5. Pipette 1µl of control DNA into the 2ml tube: Pipette 1µl of 10pg/µl control into your control transformation. Gently pipette up and down a few times. Keep all tubes on ice.
6. Close 1.5ml tubes, incubate on ice for 30 min: Tubes may be gently agitated/flicked to mix solution but return to ice immediately.
7. Heat shock tubes at 42°C for 45 sec: 1.5ml tubes should be in a floating foam tube rack. Place in a water bath to ensure the bottoms of the tubes are submerged. Timing is critical.
8. Incubate on ice for 5 min: Return transformation tubes to an ice bucket.
9. Pipette 950µl SOC media to each transformation: SOC should be stored at 4°C, but can be warmed to room temperature before use. Check for contamination.
10. Incubate at 37°C for 1 hour, shaking at 200-300 rpm.
11. Pipette 100µL of each transformation onto Petri plates Spread with a sterilized spreader or glass beads immediately. This helps ensure that you will be able to pick out a single colony.
12. Spin down cells at 6800g for 3 mins and discard 800µL of the supernatant.
13. Resuspend the cells in the remaining 100µL, and pipette each transformation onto Petri plates
14. Spread with a sterilized spreader or glass beads immediately. This increases the chance of getting colonies from lower concentration DNA samples.
15. Incubate transformations overnight (14-18 hr) at 37°C: Incubate the plates upside down (agar side up). If incubated for too long, colonies may overgrow and the antibiotics may start to break down; un-transformed cells will begin to grow.
1. Inoculate 50 ml of bacterial culture in appropriate selective media and incubate on a shaker at 250 rpm at 37℃ overnight
2. Next day, transfer the bacterial culture into a 50 ml tube and spin at 4000xg at 4℃ for 10 minutes.
3. Discard the supernatant and resuspend the pellet in 2 ml of resuspension buffer with 50 ul/ml RNase (ThermoFisher #EN053) freshly added.
4. Add 2 ml of lysis buffer to the bacterial suspension and invert the tube 3-4 times.
5. Incubate it at room temperature for 3 minutes.
6. Add 2 ml of neutralization buffer and invert the tube 3-4 times.
7. Quickly distribute the bacterial lysate into 1.5ml centrifuge tubes (approx. 4 tubes) by pouring, not pipetting.
8. Centrifuge at room temperature at 13,200xg for 10 minutes.
9. Collect supernatants in 15 ml tube and discard pellets.
10. Add 1x volume of 96% ethanol (approx. 5 ml) into the supernatant and mix it thoroughly for 5 seconds.
11. Load the sample-ethanol mix onto 5 spin-columns in three sequential (approx. 700 ul) aliquots.
12. Spin the column for 30 seconds at 13,200xg after the addition of each aliquot.
13. After each spin discard the flow through and repeat the steps till the entire sample passed through the spin columns.
14. Wash the columns 2 times with 500ul of wash buffer.
15. After each wash spin them at 13,200xg at room temperature for 30 seconds.
16. Discard the flow through.
17. Centrifuge the empty columns one more time for 1.5 minutes to remove any residue buffer.
18. After this, discard old collection tube and put the column into a new tube.
19. Add 30-35 ul of elution buffer to the column and incubate for 2 minutes.
20. Spin at 13,200xg at room temperature for 2 minutes.
21. Combine the eluted DNA from all columns in one tube (approx. 175ul).
22. After measuring the concentration store the samples at -20°C.
MEASURING THE CONCENTRATION (NANODROP)
1. Rinse Nanodrop with an ethanol and Kimtech paper towels.
2. Launch ND-8000 V2.2.1..
3. Make sure that pedestals are clean.
4. Blank with 2ul of Nuclease-Free water (work very accurately!).
5. Blank with 2ul of buffer (Elution buffer after miniprep and gel extraction).
6. Load 1ul of DNA (another person should mix the sample via pipetting up and down prior that: not vigorously!). Work very fast! Close the cover.
7. Choose wells and run.
8. Clean with ethanol and Kimtech paper towels after each use.
9. Place the cover back on.
DNA GEL ELECTROPHORESIS
1. Prepare 1% agarose gel by adding 1g of agarose powder into 100ml of 1X TAE buffer.
2. Heat it till agarose is completely dissolved (Do not boil!).
3. Add SYBR Safe as required (see the tube) in dark room.
4. Prepare the mould by wrapping it with aluminum foil.
5. Pour the solution into the mould and make sure there are no bubbles.
6. Allow the solution to set (approx. 15-20 minutes).
7. Remove comb and load ladder and samples (5ul of DNA and 1 ul of Loading Dye for 6x dye).
8. Run the gel at 100V.
was performed according to the QIAquick® Gel Extraction Kit
1. Add nuclease-free water.
2. Add buffer 5µl.
3. Add DNA up to 1000 ng.
4. Add 10 units of restriction enzyme.
5. Incubate for 30 minutes at 37°C.
6. Inactivate enzyme at 80°C for 20 minutes.
POLYMERASE CHAIN REACTION (PCR)
1. Add nuclease-free water
2.Combine 100 ng of vector and 3 fold molar excess of insert.
3. Add 5 µl ligase buffer Add 1 µl ligation enzyme
carried out by NEB protocols for Phusion Master Mix solution what we found as the best possible protocol for colony PCR. The general protocol is here.
PCR OF LIQUID CULTURE
1.Take 250 ul of culture, centrifuge at max speed for 10 min at 4℃.
2. Quickly remove supernatant and make sure it does not contain parts of the cells pellet.
3. Resuspend cell pellet in 20 ul of 0.2% Triton. Heat the tubes with cells at 98°C for 10 min.
4. Centrifuge at 14 000 g and take up the supernatant.
5. Extract the supernatant with hexane and remove the lower aqueous layer into separate tubes. This layer contains DNA. Make sure you do not take up hexane.
6. Set up PCR reaction using 3ul of DNA. Use standard reaction protocol for Phusion Master Mix with primers for SQR.
was conducted in accordance to Quan & Tian, 2011, “Circular polymerase extension cloning for high-throughput cloning of complex and combinatorial DNA libraries”. Click here for the protocol.
Na2S MEASUREMENT ASSAY
According to the information given by the company that we received our Na2S from, our sample contained at least 60% of pure Na2S.
1. Prepare a 100 mM stock solution of Na2S and bring it to pH 7.
2. This stock is diluted to 500 uM, 1 mM and 2 mM and pH is adjusted to 12 using NaOH (2M).
3. Measure the OD750 of cyanobacteria that you are willing to use (0.6 < OD750< 0.8). The OD750 of genetically modified and wild-type cyanobacteria is expected to be similar.
4. Prepare 10 mL of cyanobacteria with according concentration of Na2S.
5. Immediately after adding Na2S, take 1 mL of cells, centrifuge them for 2 mins at the highest speed and take supernatant.
6. This supernatant is measured at NanoDrop (Uv-Vis) at 230 nm. This measurement is taken before adjustment of the supernatant and nanodrop should be blanked with the normal BG-11 (pH=7).
7. The pH of the supernatant is adjusted to 12 and the measurement is repeated once again with the blank of BG-11 of the corresponding pH(pH=12).
8. Measure just BG-11+Na2S as a control with the pH 7 and 12 with corresponding blanks.
9. Repeat steps 6-7 after 5, 10, 15, 30, 60, 90 and 120 minutes.
Na2S SURVIVAL ASSAY
1. Fill 5 Petri Dish with 6 ml of liquid genetically modified and wild type strain cyanobacteria culture.
2. Add 150 uM, 250 uM, 500 uM, 1 mM, 2mM and 5mM of Na2S into the Petri Dishes for each of the genetically modified and original cyanobacteria cultures.
3. Check the color of samples during 2-4 days.
OIL SURVIVAL ASSAY
1. Make 0.1%, 0.5% and 1% oil solutions in 10 ml of liquid genetically modified and wild-type strain cyanobacteria cultures (It can be calculated given the oil density).
2. Check the OD and the color of samples for 2-4 days.