Cell Culture Protocols
Bacterial Cell Cloning / Inoculation of Cells
Entire procedure is done under flame
- For small test tubes add:
- 5mL LB media
- 500µL 20% glucose (final conc. 0.2%)
- 7.35µL of CM (final conc. 50µg/mL, working stock: 34 mg/mL)
- Add the cells
- If from glycerol stock, stab with pipette tip.
- If from LB-agar plate, pick the colony (mark it on the cap) and stab the colony
- Incubate on shaker overnight
Bacterial Cell Culture Solutions
- For 1L of LB media add:
- 10g Tryptone
- 10g NaCl
- 5g Yeast extract
- Fill to 1L with DI water
- Autoclave at P6 setting
LB-Agar Plate media
- For 1L of LB-Agar media, mix:
- 960mL water (860 mL if adding glucose)
- 10g Tryptone
- 10g NaCl
- 5g Yeast extract
- 15g Agar (add agar last)
- Autoclave (liquid option): 30 min. sterilization / 20 min drying
- Cool to 50˚C in water bath
- Next three steps are under flame
- (if adding glucose: add 100mL of 20% glucose, final conc. 2% glucose)
- Add antibiotics to the specified final conc:
- Chloramphenicol (CM): 50 µg/mL
- Ampicillin: 100 µg/mL
- Tetracycline: 50 µg/mL
- Kan: 50 µg/mL
- Pour onto the petri dishes
- Mark petri dishes using the following key:
- Ampicillin: one black line
- Chloramphenicol: two black lines
- Kan: one blue line
- Glucose: one red line
Chemically Competent Cells Transformation
This protocol uses NEB Chemically Competent Cells (ie Turbo Competent E. coli High Efficiency)
- Thaw chemically competent cells on ice.
- Transfer 50 µl of competent cells to a 1.5 ml microcentrifuge tube (if necessary).
- Add 2 µl of assembled product to NEB competent cells.
- Mix gently by pipetting up and down or flicking the tube 4-5 times. Do not vortex. Place the mixture on ice for 30 minutes. Do not mix.
- Heat shock at 42°C for 30 seconds. Do not mix.
- Transfer tubes on ice for 2 minutes.
- Add 950 µl of room temperature SOC media to tubes.
- Place the tube at 37°C for 60 minutes. Shake vigorously (250 rpm) or rotate.
- Warm selection plates to 37°C.
- Spread 100 µl of the cells onto the plates with appropriate antibiotics. Use amp plates for positive control sample.
- Incubate plates overnight at 37°C.
For overnight cell cultures 1~5mL of E. Coli in LB medium:
- Transfer cell culture into centrifuge tubes
- Centrifuge for 5 min. at 4,000 rpm
- Discard the supernatant
- Resuspend pelleted bacterial cells in 250 µl Buffer P1 and transfer to a microcentrifuge tube. Ensure that RNase A has been added to Buffer P1. No cell clumps should be visible after resuspension of the pellet.
- Add 250 µl Buffer P2 and mix thoroughly by inverting the tube 4–6 times. Do not vortex. Continue inverting the tube until the solution becomes viscous + slightly clear blue. Do not allow lysis reaction to proceed for more than 5 min.
- Add 350 µl Buffer N3 and mix immediately + thoroughly by 4-6x inverting the tube. The solution should become cloudy.
- Centrifuge for 10 min at 13,000 rpm (~17,900 x g) in a table-top microcentrifuge.
- Apply the supernatants from the previous step to the QIAprep spin column by decanting or pipetting.
- Centrifuge for 60 s. Discard the flow-through.
- Wash QIAprep spin column by adding 750µL Buffer PE and centrifuging for 60 s.
- Discard the flow-through, and centrifuge for an additional 1 min to remove residual wash buffer.
- Place the QIAprep column in a clean 1.5 ml microcentrifuge tube. To elute DNA, add 50 µl Buffer EB (10 mM Tris·Cl, pH 8.5) or water to the center of each QIAprep spin column, let stand for 1 min, and centrifuge for 1 min.
- In CryoTube vials, add:
- 250 µL of 50% glycerol
- 250 µL of cell culture
- Store in -80˚C freezer
BIO-RAD Protein Gel
- Loading dye + DTT (if not present): in 1.5mL microcentrifuge tube, make (loading dye+DTT) by:
- Get loading dye (Laemmli Sampling Buffer in 203-14A middle drawer under ``Western Blotting Supplies")
- Get DTT (in the biology section of the desiccator in the fridge--which is on the second shelf from bottom right side).
- Mix 54mg of DTT in 950µL of loading dye.
- Determine amount of protein and buffer in each lane according to following table: Note that the protein amount CANNOT exceed 8µg per lane for 10 lane gel, and 6µg per lane for 15 lane gel. Get 1.5mL microtubes as many as needed.
- Heat the tubes for 5-10 min, and get the ladder (marked "P" at the top in the Alaska fridge, in the box "Joshi Gel ladders/dyes") out to thaw.
- While the tubes are being heated, prepare the gel:
- Take the green cap off the wells and clean the wells with water. Repeat washing about three times. Take the tape off bottom.
- Assemble the cassette with two gels or (one gel + a gel-wall). Note that the wells should face inwards.
- Put the cassette into the gel box and pour NEW Tris-gly-SDS buffer into the cassette past. The buffer level should be past the wells. Check for any leaking. If there is a leak, reassemble the cassette and try again.
- Set up the voltage machine to the desired voltage and time. For a fast run, 190V with 35 minutes could work. For slow and cleaner result, 90V with 50min could work.
- Now get the heated samples, and the ladder. Add 10µL of ladder for 15 lane gels, 15µL of ladder for 10 lane gels. Add 15µL of the sample for 15 lane gels, 30µL for of the 10 lane gels.
- Fill the box to the 2-gel mark (or 4-gel mark if running more than 3 gels) or more with OLD Tris-gly-SDS buffer. Put the cap on and hit the run button. Check for bubbles rising.
Coomassie Staining Protocol
\item Dump the Tris-gly-SDS buffer into the ``used" bottle.
\item Disassemble the cassette, set the gel down on a paper towel, and rinse the rest.
\item Fetch a box to put the gel in. Crack the gel open on all four corners and retrieve the gel; be careful not the rip, and if sticky apply water. Put it in the box.
\item Check the staining solution (50\% methanol, 40\% water, 10\% acetic acid, 0.25mg per 100mL) in the fume-hood in the back corridor.
\item If the caps have been open, or the solution is out, make a new one (500mL) by:
\item 50\% of total volume is methanol (is in the flammable cabinet in the chemistry room). 40\% of total volume is water. Use plastic graduated cylinder for these.
\item 10\% of total volume is acetic acid (is in the acid cabinet). BE CAUTIOUS WHEN WALKING CORNERS, and switches gloves after touching acid. Use GLASS graduated cylinder for this.
\item measure out and add the Coomassie to appropriate amount (0.25mg$\times$ total volume). It is located in the ``Bahamas"
\item NOTE: methanol waste must go to the special waste been under bench (bench with the pH meter). If the waste bin is full, call the number.
\item Pour the Coomassie stain into the box.
- Set up plate reader
- Plate should be read at 405 nm
- Shake plate for 2-3 s before reading
- Calculate concentrations and amounts needed to reach a total reaction volume of 150 μL
- Possible conditions to vary
- Ca amount
- EGTA amount (removes Ca)
- Peptide type and concentration
- Other small molecule interactors
- Possible conditions to vary
- Dilute peptides down to working stock concentration (1.25 μM) with 20 mM HEPES buffer (should be room temp)
- Dilute working stock of peptides into multiple tubes to create range of concentrations required by assay
- Note that only 50 μL of the 150 μL well volume will be the peptide, so any concentration will be further diluted by 3x
- Usually no peptide is over 500 nM final concentration in well
- Dilute BlaCaM stock with 20 mM HEPES buffer
- Note that only 20 μL of 150 μL well volume is BlaCaM stock, so final concentration will be 2/15 of diluted stock
- Final concentration of BlaCaM in well should be 125 nM
- Add CaCl2 to 20 mM HEPES to create 10mM Ca concentration
- Plate mixtures in order onto mixing plate (buffer, CaCl2, EGTA, peptides, other molecules, BlaCaM)
- Must use non-binding plates (Corning 3641) for both mixing and assay plates to prevent binding of peptides and BlaCaM to plate
- 75.5 μL 20 mM HEPES + 10 mM Ca to all wells (5 mM final concentration)
- 50 μL 20 mM HEPES (no Ca) to BlaCaM only controls
- 70 μL 20 mM HEPES (no Ca) to blank (no BlaCaM, no peptide) controls
- 50 μL peptide to all but control wells
- Each concentration of peptide should be assayed at least three times
- 20 μL BlaCaM to all wells but blank controls
- Mix each well and incubate for 60 min on shaker (22-24°C)
- At 50 min prepare assay plate by adding 2-3 μL CENTA substrate
- Aim for final CENTA concentration of ~150 μM
- Add 97-98 μL of incubated mixture to assay plate and mix well
- Creates a total assay plate volume of 100 μL
- If possible, use Liquidator pipette system so all 96 wells can be transferred at once
- Immediately take read in plate reader and take reads every 2-3 min for 20 min
GLucCaM expression in NEB Turbo E. coli.
- Prepare 5-10mL starter culture from a fresh colony or glycerol stock in LB media supplemented with 50µg/mL CM and 0.2% glucose. Incubate at 37˚C overnight in shaker (225 rpm).
- Next day start 250mL. In 1L baffled flask, add:
- 250mL LB
- 367.5µL of CM (final conc 50µg/mL)
- 2.5mL of the cell culture grown overnight (final 1:100 dilution)
- NO GLUCOSE
- Shake in 37˚C at 225 rpm for about 3 hours to an O.D. of ~0.6
- Induce cells with 0.4mM IPTG and place immediately into an 18˚C incubator for expression. Incubate for 14-18 hours (overnight)
- Spin down cells in large floor centrifuge for 20 mins at 3000g and 4˚C
- Save cells in -20˚C freezer until ready for purification. (OR WHO KNOWS)
Agilent Herculase II PCR Amplification
Agilent Herculase II PCR Amplification Protocol:
- In a PCR tube, mix (50 µL final volume):
- 35 µL water
- 10 µL HercII Buffer (5X)
- 0.5 µL dNTP (25mM each, 100mM total final)
- 1.25 µL forward primers (10 µM)
- 1.25 µL reverse primers (10 µM)
- 1 µL Template DNA (diluted to 10 ng/µL)
- 1 µL HercII DNA polymerase
- Give the tubes a brief spin
- Thermocycling conditions (see diagram below):
- Step 1: 95˚C, 2:00
- Step 2 X30
- Phase 1: 95˚C, 0:15
- Phase 2: (Tm - 5)˚C, 0:20
- Phase 3: 72˚C, 0:30 per 1kb
- Step 3: 72˚C, 3:00
- First dilution: dilute to 100 µM (shortcut: amount_in_nMoles X10 µL)
- Second dilution: dilute to 10 µM
For 2-3 fragment assembly
- Set up the following on ice in PCR tube (total volume 20µL):
- 0.02–0.5 pmols of fragments: 50-100ng of vectors with 2-3 fold of excess inserts.
- 10 µL of Gibson Assembly Master Mix (2X)
- Fill to 20µL with DI water
- Incubate in a thermocycler at 50˚C for 60 min.
- Wipe the lower and upper measurement pedestals with 2µL DI water
- Measure blank by pipetting 2µL of sample solvent (usually water or DNA elution buffer) on the lower pedestal
- Measure conc. of samples by 2µL on the lower pedestal
ZYMO DNA Clean & Concentrator
ZYMO DNA Clean & Concentrator - 5 Kit was used to purify DNA from PCR.
- Add 2-7 Volumes of DNA Binding Buffer to each volume of DNA sample.
- For plasmid or genomic DNA 2kb, add 2 volumes.
- For PCR or short DNA fragments, add 5 volumes.
- For ssDNA purification, add 7 volumes
- Load the mixture into a Zymo-Spin Column in a Collection Tube.
- Centrifuge at full speed (≥10,000 x g) for 30 seconds. Discard flow through.
- Add 200 µl of DNA Wash Buffer to the column and centrifuge for 30 seconds.
- Repeat previous step, this time centrifuging for 1 minute.
- Place the Zymo-Spin Column into a new 1.5 ml tube. Add ≥6 µl of DNA Elution Buffer or water directly to the column matrix and spin (30 seconds) to elute the DNA.
GeneWiz Sequencing Prep
The following protocol should be followed to prepare DNA for submission to GeneWiz for sequencing.
- Complete online order form.
- Label PCR tubes with the appropriate names form the order form print out.
- Concentrate template DNA according to the table below.
- Mix 10 µl of DNA template with 5 µl of primer.
- Attach PCR tubes to order form and submit GeneWiz mailbox.