Team:WashU StLouis/Protocols

Preparation of Materials E. coli Transformation Yeast Transformation DNA Separation and Preparation DNA Manipulation

Preparation of Materials

Fill a container to about 70% of its volume with distilled water (assuming the container volume is the total volume of media intended to be made).
Add 10 g of tryptone.
Add 5 g of yeast extract.
Add 10 g of NaCl.
Fill up remaining volume with distilled water.
Stir until all LB Broth is dissolved.
Autoclave at 121 ˚C for 100 minutes.
Adjust pH to 7.0 with NaOH.

Fill a container to about 60-70% of its volume with distilled water (assuming the container volume is the total volume of media intended to be made).
Add 10 g of tryptone.
Add 5 g of yeast extract.
Add 10 g of NaCl.
Add 15 g/L of Agar.
Fill up remaining volume with distilled water.
Stir until all solid is dissolved.
Autoclave at 121 ˚C for 100 minutes.

Heat previously prepared LB agar so that it has completely melted.
Allow agar to cool.
Add enough antibiotic solution to the LB agar so that there is 1 microliter of antibiotic solution per 1 mL of LB agar.
Add about 20 mL of the LB agar + antibiotic mixture into each empty plate, and make sure that the entire plate is covered.
Let the mixture cool and harden and then store the plates at 4 ˚C.

Fill a container to about 400 mL.
Add 6 g of yeast extract.
Add 12 g of peptone.
Fill up remaining volume to 600 mL with distilled water.
Autoclave at 121 ˚C for 100 minutes.
After cooling media to less than 50 ˚C, add 12 g of filter sterilized glucose.

Fill a container to about 400 mL.
Add 6 g of yeast extract.
Add 12 g of peptone.
Add 60 mg of adenine hemisulphate.
Fill up remaining volume to 600 mL with distilled water.
Autoclave at 121 ˚C for 100 minutes.
After cooling media to less than 50 ˚C, add 12 g of filter sterilized glucose.

Fill a container to about 400 mL.
Add 6 g of yeast extract.
Add 12 g of peptone.
Add 20 g of agar.
Fill up remaining volume to 600 mL with distilled water.
Autoclave at 121 ˚C for 100 minutes.
After cooling media to less than 50 ˚C, add 12 g of filter sterilized glucose.

Fill a container with 800 mL of distilled water.
Add 80 g of NaCl.
Add 2 g of KCl.
Add 14.4 g of Na₂HPO₄.
Add 2.4 g of KH₂PO₄.
Fill to 1 L with distilled water.
Stir until all solid is dissolved.
Autoclave at 121 ˚C for 100 minutes.

Add 30 g of Na₂HPO₄
Add 15 g of Na₂HPO₄
Add 5 g of NH₄Cl
Add 2.5 g of NaCl
Add 15 mg of CaCl₂
Add 1 L of of distilled water

Add 5 g of PEG 8000.
Add 0.30 g of MgCl₂*6H₂O.
Add 2.5 mL of DMSO.
Fill to 50 mL with LB media.
Filter sterilize using a 0.22 µL filter.
Store at 4 or -20 ˚C.

Fill a container with 700 mL of distilled water.
Add 20 g of bacto-tryptone.
Add 5 g of bacto-yeast extract.
Add 0.584 g of NaCl.
Add 0.186 g of KCl.
Add 2.4 g of MgSO₄.
Fill to 1 L with distilled water.
Stir until all solid is dissolved.
Autoclave at 121 ˚C for 100 minutes.
After cooling media to less than 50 ˚C, add 3.603 g of glucose.
Prior to use, adjust pH to 7.5 by adding approximately 25 mL of 1M NaOH.
Filter sterilize the media before use.

Add 750 μL 50% glycerol to cryotube.
Add 750 μL liquid culture to cryotube.
Store in -80°C freezer.

Add 5 mL of LB into a culture tube.
Use a pipette tip or inoculation loop to pick a single colony.
Dip the tip or loop into the LB media.
Incubate at 37 ˚C.

Measure 30 mg of solid Chloramphenicol per mL of antibiotic intended to be made.
Dissolve the Chloramphenicol in 100% EtOH and fill to the volume intended.
Filter the Chloramphenicol solution through a syringe with an appropriate filtration tip.
Aliquot the solution into microcentrifuge tubes and store at -20°C.

Measure 100 mg of solid Ampicillin per mL of antibiotic intended to be made.
Dissolve the Ampicillin in 100% EtOH and fill to the volume intended.
Filter the Ampicillin solution through a syringe with an appropriate filtration tip.
Aliquot the solution into microcentrifuge tubes and store at -20°C.

Measure 50 mg of solid Kanamycin per mL of antibiotic intended to be made.
Dissolve the Kanamycin in 100% EtOH and fill to the volume intended.
Filter the Kanamycin solution through a syringe with an appropriate filtration tip.
Aliquot the solution into microcentrifuge tubes and store at -20°C.

Spin down 5 mL of cells at an OD between 5-15 for 5 min at 10,000 g.
Take 2 mL of cold .9% filtered NaCl and resuspend the cells on ice.
Recentrifuge.
Resuspend in approximately 1-1.5 mL of cold 30% chloroform, 70% MeOH solution using glass vials.
Shake for 4 hours at 4°C and vortex every hour for 15 sec.
Add 0.5 mL of ddH₂O and spin gently for 2 min.
Take upper aqueous fraction, store in glass gc vials.
Repeat step 6 and store in gc vials.

Lypholyze overnight.
Redissolve dry sample in 100 uL of N-methyl-N-(trimethylsiyl) trifluoroacetamide.
Derivatize at 70°C for 1 hr.
Run on GC-MS, along with standards and spiked samples.

E. coli Transformation

Materials:

LB Media
Pre-chilled TSS Buffer
Pre-chilled 1 mL Eppendorf tubes
Pre-chilled Falcon tubes

Grow a 5 mL overnight culture in LB media for 16-20 hours.
Dilute the overnight culture by 100 fold volume of LB media in a new culture tube (100 uL culture in 10 mL LB).
Incubate at 37 ˚C with continuous shaking.
Monitor the growth of cells and stop when OD600 reaches 0.3-0.35.
Make sure the centrifuge is fully cooled and spin down the cells at 2000 g at 4 ˚C in Falcon tubes for 10 minutes.
Remove the supernatant.
Resuspend the cells in one tenth the volume of the original culture, after taking into account the loss of volume from monitoring, of pre-chilled TSS buffer while keeping cells on ice (1 mL TSS).
Aliquot 100 µL cells per tube while keeping cells on ice.
Freeze the cells in liquid nitrogen and store at -80 ˚C.

Thaw 100 µL of competent cells on ice.
Add 1 µL of DNA (plasmids) to the thawed competent cells.
Incubate the cells and DNA on ice for 30 minutes.
Place the tubes with the cells and DNA in a 42 ˚C bath for 35-45 seconds.
Leave cells on ice for about 5 minutes.
Warm SOC media to room temperature and filter sterilize.
Add six times the cell quantity µL of SOC media to the cells.
Incubate at 37 ˚C for 90 minutes, shaking at 50 rpm while the tubes rest sideways.
Warm plates to room temperature for 90 minutes.
Pipette 50-200 µL of each transformation onto petri plates.
Incubate transformations overnight (14-18 hours) at 37°C with plates upside down.

Grow an overnight culture for 16-20 hours at 37°C.
Dilute overnight culture 100-fold into fresh LB medium (2 mL in 200 mL).
Grow cells in 37°C until OD600 reaches 0.3-0.35 (Measure OD every hour, then every 15-20 minutes once OD reaches 0.2).
Immediately put the cells on ice and chill cells for 10 minutes (swirl/invert occasionally to ensure even cooling) and pre-chill centrifuge to 4C.
Split the cells into four 50 mL conical tubes and spin down at 2000 g for 10 minutes at 4˚C.
Decant the supernatant.
Add 40 mL of ice cold, sterile ddH2O to each 50 mL conical tube and pipette cells back into solution with a serological pipet.
Spin at 2000 g for 10 minutes at 4˚C.
Decant the supernatant.
Add 20 mL of ice cold sterile ddH2O to each 50 mL conical tube and pipette cells back into solution with a serological pipet.
Spin at 2000 g for 10 minutes at 4˚C.
Decant the supernatant.
Add 4 mL of ice cold sterile 10% glycerol to each 50 mL conical tube and pipette cells back into solution by pipetting and combine the four suspensions into a single 50 mL conical tube.
Spin at 2000g for 10 minutes at 4˚C.
Decant the supernatant.
Add 400 uL of ice cold sterile 10% glycerol to the 15 mL conical tube. Resuspend with a 1 mL pipette.
Label and pre-chill 1.5 mL centrifuge tubes.
Aliquot suspended cells in 20-50 µL aliquots (make sure not to touch cells with warm fingers).
Store aliquots in -80˚C.

Combine 20 µL of cells and 1-500 ng of plasmid (or ligation), making sure DNA volume does not exceed 10% of the volume of competent cells (maximum of 5 uL of DNA in 50 uL of cells).
Immediately pipette cell and DNA solution into a 1 mm electroporation cuvette (pre-chilled on ice).
Tap the electroporation cuvette on the bench to ensure suspension has fallen to the bottom and has spread through the entire width of the cuvette.
Gently pat/wipe down the water on the outside of the electroporation cuvette with a kimwipe.
Pulse the electroporation cuvette using protocol Ec1.
Resuspend cells in 9 parts LB media, aspirate up and down to mix the suspension, and move the diluted suspension to a 1.5 mL centrifuge tube (tilt cuvette and pipette slightly to the side to ensure all of the solution is captured.
Incubate at 37 ˚C for 40-60 minutes, shaking at 250 rpm.
Add LB media to a final volume of 1 mL.
Plate 100 µL of cells.
Spin down the remaining 900 µL of cells at a maximum of 6000 g for 3 minutes.
Remove about 700 µL of the supernatant, resuspend, and plate the resuspended cells.
Incubate plates upside down at 37 ˚C.

Thaw cells on ice. Label one 1.5 mL microcentrifuge tube for each transformation and prechill (do three per plasmid concentration and average).
Spin down DNA tubes from competent cells kit at 8,000 to 10,000 rpm for 20-30 seconds to collect all DNA at bottom.
Pipet 50 µL of competent cells into each tube while keeping cells on ice.
Pipet 1 µL of DNA into each tube while keeping cells on ice (do not exceed 10% of the competent cell volume).
Incubate on ice for 30 minutes and preheat water bath to 42 ˚C.
Heat shock cells for 35-45 seconds (keep lids of tubes above water and and ice close by).
Immediately transfer tubes back to ice and incubate for 1-2 minutes.
Warm SOC to room temperature and filter sterilize
Add 350 µL of SOC media per tube and incubate for 90 minutes at 37 ˚C shaking at 200-300 rpm while the tubes are sideways.
Warm plates to room temperature.
Pipet 50-200 µL from each tube onto agar + CM plate and spread evenly.
Incubate plates overnight at 37 ˚C.
Count number of colonies and use equation to calculate your competent cell efficiency (average triplicates for each sample).
Efficiency (in cfu/µg) = [colonies on plate (cfu) / Amount of DNA plated (ng)] x 1000 (ng/µg)

Yeast Transformation

Take plate containing desired strain from the 4°C room and pick a medium size colony.
Inoculate the colony in 5 mL of YPAD.
Add 5 uL of Kanamycin to inoculum.
Place tube in 30°C shaker overnight (200 rpm)
Prepare 25 mL of YPAD in a flask and place in the 30°C shaker overnight.
Take the small falcon tube from the 30°C shaker and prepare for OD₆₀₀ measurements.
Prepare 1 cuvette with 1000 uL of YPAD (blank) and 1 cuvette with 40 uL of yeast culture in 960 uL of YPAD.
Blank the spectrophotometer using the pure YPAD sample then record the OD of the sample of interest.
Add the appropriate amount of yeast culture to the 25 mL or 50 mL flask such that the resulting OD of the flask is 0.2.
After adding appropriate volume of culture (Y) to pre-warmed 25 mL or 50 mL flask of YPAD, incubate the mixture at 30°C on a shaker at 200 rpm until it is equivalent to 2x10⁷ cells/mL (OD₆₀₀ = 0.8). This will take approximately 3 to 5 hours, depending on the strain of yeast.
Harvest the culture in a sterile 50 mL centrifuge tube at 4000 rpm for 5 min.
Pour off the supernatant and resuspend the cells in 25 mL of ddH20 and centrifuge again.
Pour off the supernatant water and resuspend the cells in 1.0 mL of 100 mM LiAc and transfer the suspension to a 1.5 mL microcentrifuge tube.
Pellet the cells at 6000 rpm for 15 sec and remove the LiAc with a micropipette.
Resuspend the cells to a final volume of 500 uL (2x10⁹ cells/mL)- about 400 uL of 100 mM LiAc.
Boil SS-DNA for 5 min and quickly chill in ice water.
Vortex the cell suspension and pipette 100 uL samples into microcentrifugte tubes. Pellet the cells and remove the LiAc with a micropipette.
The basic transformation mixture consists of the following (add in listed order):

240 uL PEG 3350 (50% w/v)
36 uL 1.0 M LiAc
50 uL SS-DNA (2.0 mg/mL)
X uL (200 ng) DNA vector
34-X uL sterile ddH₂O

Vortex each tube vigorously until the cell pellet has been completely mixed.
Heat shock in a 42°C water bath for 50 min.
Microcentrifuge at 6000-8000 rpm for 15 sec and remove the transformation mix supernatant with a micropipette.
Pipette 1000 uL of sterile ddH₂O into each tube and resuspend the pellet by pipetting it up and down gently.
Plate onto solid media.
Incubate plates for 2-4 days at 30°C to recover transformed cells.

DNA Separation and Preparation

Incubate a cell culture overnight at 37˚C.
Transfer the culture to 50 mL Falcon tubes.
Centrifuge the tubes at 3000 RPM for 2 minutes and decant the supernatant.
Resuspend the cell pellets in 250 µL of resuspension solution.
Add 250 µL of lysis buffer.
Add 350 µL of neutralization solution and mix immediately.
Centrifuge the tubes at 10,000 RPM for 15 minutes.
Transfer to spin columns with microcentrifuges attached.
Centrifuge for 1 minute at 10,000 RPM and discard the flow-through.
Add 500 microliters of wash solution into the spin column.
Centrifuge for 30-60 seconds at 10,000 RPM and discard the flow-through.
Repeat the previous two steps.
Centrifuge for 1 minute at 10,000 RPM.
Transfer the spin columns onto new 1.5 mL microcentrifuge tubes and add 50 µL of elution buffer.
Incubate for 2 minutes at room temperature.
Centrifuge for 2 minutes at 10,000 RPM.
Discard the spin column and store the DNA solution in the microcentrifuge tube at -20 ˚C.

Mix 75 mL of TAE (Tris-acetate-EDTA) buffer with 0.75 g of agarose so that the mass concentration is 1% (need lower concentration for larger bands and a higher concentration for smaller bands)
Microwave the solution until all the agarose dissolves in the TAE buffer.
Let the solution cool to 55 ˚C, or until the flask can be held by a gloved hand.
Add and mix 75 µL of SYBRsafe into the TAE-agarose solution and stir gently.
Pour the solution into the gel cassette.
Place a comb onto the the cassette to make the wells.
Let the gel solidify for 30 minutes.
Combine 10 µL of the digestion with 2 µL of 6x loading dye.
Place the gel with the cassette into the electrophoresis box and add TAE buffer until it covers the entire gel.
Remove the comb to expose the empty wells.
Add 5 - 6 µL of DNA ladder to the first well.
Pipette the digestion and dye mixture into the wells.
Set the voltage to 120V and run the gel until the dye runs approximately three-fourths of the way down the gel (about 30-45 minutes).
Using the proper protection, view the bands with UV light, and cut out the wanted bands which can be stored at -20 ˚C in a microcentrifuge tube.

Determine the mass of each gel fragment by weighing it and subtracting the mass of an empty microcentrifuge tube.
Add Buffer QG so that there is 300 µL of buffer for every 100 mg of agarose gel.
Incubate at 50 ˚C for 10 minutes or until the gel has dissolved and mix by vortexing every 2-3 minutes.
Check that the color of the mixture is yellow and similar to that of Buffer QG, adjusting pH if necessary.
Add 100 µL of isopropanol for 100 mg of agarose gel and mix.
Place a spin column in a 2 mL collection tube.
To bind DNA, add the sample to the spin column and centrifuge for a minute at over 10,000 g.
Discard flow-through and place spin column back in the same collection tube.
Add 0.5 mL of Buffer QG and centrifuge for a minute at over 10,000 g.
To wash, add 0.75 mL of Buffer PE, with ethanol added, to the spin column and centrifuge for a minute at over 10,000 g.
Discard flow-through and centrifuge for an additional minute at over 10,000 g.
Place spin column into a clean 1.5 mL microcentrifuge tube.
To elute DNA, add 50 µL of Buffer EB to the spin column and centrifuge for a minute at maximum speed. (For increased DNA concentration, add 30 µL of elution buffer, let column stand for 1 min, and then centrifuge for a minute at maximum speed)
Discard the spin column and store DNA at -20 ˚C.

Materials:

Minipreped samples A (sequence that comes first) and B
Linearized plasmid backbone with a different antibiotic resistance than plasmid samples
EcoRI, Xbal, SpeI, PstI, DpnI
NEB Buffer 2
BSA
dH2O

Part 1: Digestion

Make the Enzyme Master Mix for the Plasmid Backbone (25 µL, for 5 reactions).

5 µL NEB Buffer 2
0.5 µL BSA
0.5 µL EcoRI-HF
0.5 µL PstI
0.5 µL DpnI (Used to digest any template DNA from production)
18 µL dH2O

Make the Enzyme Master Mix for the Part A (25 µL, for 5 reactions).

5 µL NEB Buffer 2
0.5 µL BSA
0.5 µL EcoRI-HF
0.5 µL SpeI
18.5 µL dH2O

Make the Enzyme Master Mix for the Part B (25 µL, for 5 reactions).

5 µL NEB Buffer 2
0.5 µL BSA
0.5 µL XbaI
0.5 µL PstI
18.5 µL dH2O

Digest Plasmid Backbone, Part A, and Part B at 37 ˚C for 30 minutes individually by adding 4 µL of DNA (25 ng/µL) and 4 µL of the corresponding Enzyme Master Mix.
Heat kill at 80 ˚C for 20 minutes.
Discard the spin column and store eluted DNA at -20°C.

Part 2: Ligation

Add 2 µL of digested Plasmid Backbone (25 ng).
Add equimolar amount of Part A fragment (< 3 µL).
Add equimolar amount of Part B (< 3 µL).
Add 1 µL T4 DNA ligase buffer.
Add 0.5 µL T4 DNA ligase.
Add water to 10 µL.
Ligate by incubating at 16 ˚C for 30 minutes.
Heat kill at 80 ˚C for 20 minutes.
Transform cells with DNA.

Prepare DNA inserts (for g blocks, resuspend according to given instructions; for plasmids, transform bacteria, allow replication, and perform miniprep).
Perform the 3A Assembly digestion and ligation protocol.
Transform chassis with plasmids and plate cells.
Incubate transformations overnight (14-18 hours) at 37°C with plates upside down.
Pick and grow an individual overnight culture in LB+antibiotic for 16-20 hours for 4-6 colonies.
Make a glycerol stock for each colony’s overnight culture.
Perform the miniprep protocol on the overnight cultures.
Perform the digestion protocol using restriction enzymes that yield a large fragment size.
Run the gel electrophoresis protocol to check for correct band size.
Grow an overnight culture in LB+antibiotic for 16-20 hours of a glycerol stock that contained the correct band size.
Perform the miniprep protocol on the overnight culture.
Sequence the isolated DNA.

DNA Manipulation

Add 1 to 5 µL of DNA solution to a microcentrifuge tub.
Add 5 µL of digestion buffer.
Add nuclease free and autoclaved H₂O until the total volume reaches 48 µL.
Add 1 µL of each of the two enzymes and pipette up and down to mix.
Incubate the tube at 37 ˚C for 2 hours.

Using the concentration of DNA for the backbone and insert, the number of base pairs in each segment of DNA, and the desired molar ratio, calculate and add the appropriate masses and volumes of insert DNA and plasmid backbones into a microcentrifuge tube.
Add 1 µL of Ligase Buffer.
Add nuclease free and autoclaved H₂O so that the total volume becomes 9 microliters.
Add 1 µL of DNA ligase.
Incubate the ligation mixture at room temperature (time will vary per ligase).
After incubation, the DNA can be transformed into competent cells or stored at -20°C.

Gently vortex and briefly centrifuge PCR Master Master Mix (2x) after thawing.
Place a thin-walled PCR tube on ice and add 25 µL of PCR Master Mix (2x), 1 µL of 25 µM forward primer, 1 µL of 25 µM reverse primer, 1 µL of 10 ng/µL template DNA, and 22 µL of nuclease free and autoclaved H₂O for a total volume of 50 µL.
Gently vortex the sample and spin down.
Perform PCR using the proper thermal cycling conditions.
Perform gel purification steps or store the PCR sample at -20 ˚C