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Team:Washington/Protocols
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Protocols
Media, Plates, and Solutions
Competent Cell Media Buffer (CCMB)
Mix the following to a 2 L container:
100 g glycerol (liquid)
10 mL x 1 M potassium acetate
11.8 g CaCl2*H2O
4 g MnCl2
2 g MgCl2
1 L of dH2O
Sterile filter or autoclave (20 min at 121 °C and 20 psi) in a 1 L bottle
Super Optimal Broth (SOB)
Mix the following to a 2 L container:
20 g Tryptone
5 g Yeast Extract
10 mL x 1 M NaCl
2.5 mL x 1 M KCl
1 L of dH2O
Sterile filter or autoclave (20 min at 121 °C and 20 psi) in a 1 L bottle
Phosphate Buffered Saline (PBS) Solution
Mix the following to a 2 L container or 1 L beaker:
8 g NaCl
1.44 g Na2HPO4
0.8 g KCl
0.24 g KH2PO4
1 L of dH2O
Buffer to pH 7.4
Sterile filter or autoclave (20 min at 121 °C and 20 psi) in a 1 L bottle
PSBF (PBS for Flow)
Mix the following in a 1 L beaker:
25 mL 20X PBS, pH 7.4
574 mL H2O
2.5 g BSA (0.5%)*
Filter and store at 4 °C
*Do not need to pH - should be at pH 7.4 like 20X PBS
Yeast Extract Peptone Dextrose (YPD)
Mix the following into 950 mL of dH2O in a 1 L bottle:
20 g peptone
10 g yeast extract
Autoclave (20 min at 121 °C and 20 psi)
Add 50 mL 40% glucose
Sterile filter into a 1 L bottle
Note: For long-term liquid media storage, do not add 40% glucose. Instead add the glucose directly into cell cultures.
Note: For YPD-plates add 24 g agar to the peptone and yeast extract before autoclaving.
Selective Dropout Media, C-Uracil and C-Histidine (C-Ura and C-His)
Synthesized by the Yeast Resource Center at the University of Washington's Department of Genome Sciences and Department of Biochemistry.
Guanidinium Hydrogen Chloride
For maximum effectiveness, final concentration should be approximately 8.5 M in PBS.
203 g guanidinium hydrogen chloride
250 mL PBS solution*
Add dilute HCl to pH 7.4
*Alternatively add slightly less than 250 mL of PBS in order to buffer the solution to the appropriate volume, then add more dH2O as necessary.
Basic Cloning
Polymerase Chain Reaction
All PCRs were done using a standard 50 μL reaction volume with GoTaq® Green Master Mix 2X purchased from PROMEGA Corporation.
Mix the following in a 0.2 mL microcentrifuge tube on ice:
25 μL GoTaq® Green Master Mix 2X
1-5 μL of 10 μM forward primer
1-5 μL of 10 μM reverse primer
<250 ng of DNA template
QS 50 μl nuclease-free H2O
Conduct the reaction in a thermocycler, adjusting anneal temperature and extension times accordingly. See your polymerase supplier protocol for more details on thermocycling.
Error-prone Polymerase Chain Reaction
Prepare 50 μL reaction:
5 μL 10X Mutazyme II Rxn Buffer
1 μL 40 mM dNTP mix (200 μM each final)
1 μL 20 μM forward primer
1 μL 20 μM reverse primer
1 μL Mutazyme II DNA polymerase (2.5 U/μL)
0.01 ng template
QS 50 μL diH2O
Program thermocycler as follows:
95 °C, 2 min
95 °C, 30 sec
XX °C*, 30 sec
72 °C, X min**
32 cycles
72 °C, 10 min
4 °C, hold
*Adjust annealing temperature according to Tm of primer.
**Adjust extension time according to the length of amplified DNA.
Note: Use 0.01 ng of template (calculate by insert and not by total plasmid).
Calculate amount of template to use as follows:
(bp for amplified region) / (bp in total plasmid) = % amplified region
(conc. of total plasmid) x (% amplified region as a decimal) = conc. of amplified region
Note: Never pipette less than 0.5 μL.
(0.01 ng of template) / (conc. of amplified region) = vol of template to add to PCR
Restriction Endonuclease Reaction (Digestion)
All restriction enzyme reactions were done using a 50 μl reaction volume. Restriction enzymes and buffers were purchased from New England Biolabs® Inc.
Mix the following in a 0.2 mL PCR tube:
1 μg of DNA
5 μL of the appropriate 10X New England Biolab® Buffer
1 μL of each restriction enzyme (add last)
QS 50μL nuclease-free H2O
Incubate the reaction for 1 hr
Heat inactive the reaction at the appropriate temperature
Note: Thaw the restriction enzyme(s) on ice to improve shelf life
Ligation
T4 DNA Ligase and Buffer were purchased from New England Biolabs® Inc.
1. Prepare the following in a 0.2 mL microcentrifuge tube:
50.0 ng vector DNA*
37.5 ng vector DNA*
2 μL 10X T4 DNA Ligase Buffer
1 μL T4 DNA Ligase
QS 20 μL diH2O
2. Incubate the reaction at room temperature for 10-30 minutes or at 16 °C overnight.
3. Heat inactivate at 65 °C for 10 minutes.
4. Chill on ice before starting a transformation reaction.
*The exact amount of DNA is dependent on the number of base pairs. In order to conduct a proper reaction consult the New England Biolab Ligation Calculator at:
http://nebiocalculator.neb.com/#!/
Escherichia coli Protocols (XL1-Blue and XL10-Gold)
Chemically Competent Cell Cultures
1. Plate 10 ul of cells onto an antibiotic free LB/Agar plate..*
2. Incubate overnight at 37OC
3. Inoculate lots of colonies into 50ml SOB-Mg media in a 250ml flask.
4. Incubate at 37 OC, 200rpm, until OD at 550nm reaches 0.3, takes about 3 hours. I do my first check at 2 hours.
5.Transfer the cell culture into a sterile 50ml Falcon tube and chill on ice for 10 minutes.
6. Pellet the cells at 2500rpm for 15 minutes at 4 OC.
7. Decant supernatant and invert tubes to remove excess culture medium.
8. Resuspend the cells in 16ml CCMB by gentle vortexing or pipetting (I shake the falcon tube).
(16ml CCMB per 50ml culture)
9. Incubate on ice for 20 minutes.
10. Centrifuge at 2500rpm for 10 minutes at 4 OC.
11. Decant supernatant and invert tubes to remove excess liquid.
12. Resuspend the cells in 4ml CCMB by gentle vortexing or pipetting.
(4ml CCMB per culture)
13. If making multiple 50ml cultures, combine all of the cells and mix before aliquoting.
14. In cold room: make 205ul aliquots, flash freeze in liquid nitrogen, and store at -80 OC.
NOTE: Make SOB-Mg media and autoclave the 250ml flasks with water in them the same day the cells are plated (step 1)
Chemically Competent Cell Transformations
1. Thaw competent E.coli cells on ice (XL1-Blue or XL10-Gold)*
2. Add 50μL of competent cells to sterile 15mL conical centrifuge tubes
3. Add 1μL (~100-200ng)* of the mini-prep to each culture tube
4. Equilibrate the cells on ice for 10 min
5. Heat shock the cells at 42C for 30-45 seconds**
6. Immediately place the cells back on ice for 3 min
7. Add 250μL LB media without antibiotics and shake at 250 rpm and 37C for 30 min
8. Spread 10μL and 290μL on an appropriate LB-antibiotic plate
9. Invert the plate and incubate at 37C overnight
*The exact amount of DNA to add depends on your cell's transformation efficiency. However, it is acceptable to add a larger amount to increase the number of transformed cells.
** Do not heat shock for an extended duration as this may damage and/or kill your cells.
Overnights
1. In a 14mL round-bottom tube, add 3-5mL of LB and an appropriate volume of antibiotic(s).
2. Swipe several individual colonies, do not collect satellites or colony clumps, with a pipette tip.
3. Swirl the colony tip in the tube, there should be no visible cell clumps.
4. Incubate and shake the tube at 37C at 250rpm for 12-16 hours and no longer than 20 hours.
DNA-Extraction and mini-preps
All DNA mini-preps were prepared using EPOCH mini-kits and following the supplied protocols.
Glycerol Stocks
1. Take 1-2mL from an overnight culture and transfer into a 1.5mL centrifuge tube.
2. Spin down the culture at 3000rpm for 3 minutes.
3. Decant the supernatant.
4. Resuspend the cells in 500μL of 40% Glycerol and 500uL of LB (no antibiotics) or water.
5. Transfer the resuspension to a cryogenic vial.
6. Store the glycerol stock at -80C.
Saccharomyces cerevisiae (PYE1 Yeast)
Chemically Competent Cell Culturing
This process take 4 days in lab with a 1 day wait for incubation.
Day 1:
1. Streak yeast cells onto a YPD plate.*
2. Invert the plate and incubate at 30C for 2 days.
Day 3:
1. Add 50mL of YPD liquid media into a 250mL baffle flask.
2. Swipe as many individual colonies as you can see into the YPD media.**
3. Incubate and shake the culture at 30C at 250rpm overnight approximately 24 hours.
Day 4:
1. Take an optical density measurement.
2. In three 250mL baffle flask add the portions of the overnight liquid culture.
3. Dilute each culture to approximately 0.4 optical density with YPD.
4. Incubate and shake the cultures at 30C at 250rpm until the optical density reaches 1.2-1.6.
5. Collect each culture into separate 50ml flat-bottomed centrifuge tubes.
6. Spin down the cells at 4000g for 5 minutes at 4C.
7. Decant the supernatant.
8. Resuspend the cells in 100mL total for all three culture of dH2O.
9. Combine the suspensions into two 50mL flat-bottomed centrifuge tubes.
10. Spin down the cells as above.
11. Decant the supernatant.
12. Resuspend each in 3mL of 100mM Lithium Acetate.
13. Transfer both cultures into a single 15mL conical centrifuge tube.
14. Spin down the cells at 3000rpm for 5 minutes.
15. Resuspend the cells in 0.75mL of 100mM Lithium Acetate, total volume is roughly 2mL.
16. Qualitatively bring up the volume to 3.5mL by adding 40% Glycerol.
17. Aliquot the cells into 1.5mL centrifuge tubes or 1.7mL cryogenic vials.***
*Streak in such a way that there are individual colonies visible on the plate without clumps or satellite colonies.
**Collect only individual visible colonies. Do not collect clumps or satellite colonies.
***The volume of aliquots depends on the number to transformations you intend to do at a time.
Chemically Competent Transformations
This protocol assumes a 50μL aliquot of yeast competent cells were made.
Furthermore, this protocol prepares enough cells for 6 yeast transformations.
1. Add the following to 50μL of yeast competent cells:
240μL of Polyethylene Glycol - 3350 (PEG-3350)
36μL of 1M Lithium Acetate
32μL of dH2O
2. Mix the mixture by gentle pipetting or vortexing.
3. Aliquot 59μL of the mixture into a 0.2mL microcentrifuge tube.
4. Add 1uL (~100-200ng) of DNA.
5. Mix the mixture by gentle pipetting or vortexing.
6. Incubate the mixture at 30C for 30 minutes.
7. Heat shock the mixture at 42C for 20 minutes.
8. Spin down the cells in a microcentrifuge for ~1 minute.
9. Decant the supernatant.
10. Resuspend the cell pellets in 200μL of dH2O.
11. Spin down the cells in a microcentrifuge for ~1 minute.
12. Resuspend the cell pellets in 200μL of dH2O.
13. Plate 50-150μL of the mixture onto an appropriate Selective Dropout Media plate.
14. Invert and incubate at 30C for 2 days.
*The exact amount of DNA depends on the transformation efficiency of your competent cells.
Overnight Culturing
1. In a 14mL round-bottomed culture tube add 1.8mL selective dropout media and 0.2mL 20% glucose.
2. Swipe 3 individually visible yeast colonies and add them to the culture tube media.
3. Incubate and shake at 37C at 250rpm for 2 days.
Note: You can also do 3mL cultures (2.7mL S.D. media and 0.3mL 20% glucose) or larger cultures just make sure to dilute the glucose from 20% to 2%.
Culture Passaging
1. In a 14mL round-bottomed culture tube add 1.8mL selective dropout media and 0.2mL 20% glucose.
2. Take 20-50μL from a previous overnight or passage culture and add it to the culture media.
3. Incubate and shake at 37C at 250rpm for 2 days.
Note: You can also do 3mL cultures (2.7mL S.D. media and 0.3mL 20% glucose) or larger cultures just make sure to dilute the glucose from 20% to 2%.
Note: The exact amount of culture that you take from a previous culture is irrelevant as long as at least 1 living cell is passaged.
Glycerol Stocks
1. Take 1-2mL from an overnight culture and transfer into a 1.5mL centrifuge tube.
2. Spin down the culture at 3000rpm for 3 minutes.
3. Decant the supernatant.
4. Resuspend the cells in 500uL of 40% glycerol and 500μL of Selective Dropout media or water.
5. Transfer the resuspension to a cryogenic vial.
6. Store the glycerol stock at -80C.
Flow Cytometry
Dilutions
1. From an overnight culture measure the optical density at 660nm by making 1:10 dilutions.
2. Take enough culture to make a 1mL aliquot with OD 0.4.
3. Spin down the aliquot in a 1.5mL centrifuge tube at 3000rpm for 3 minutes.
4. Decant the supernatant.
5. Resuspend the cell pellet in 800μL of the appropriate selective dropout media and 200μL of 20% glucose.
6. Transfer the new culture to a 14mL culture tube.
7. Incubate and shake at 30C and 250rpm for at least 6 hours (1.2-1.6 optical density).
Preparations for Analysis using C6 Accuri Flow Cytometer
1. From the dilution previously made, measure the optical density, roughly 1.2-1.6.
2. Make an aliquot of 500μL of the dilution culture in a 1.5mL centrifuge tube.
3. Spin down the aliquot at 3000rpm for 3 minutes.
4. Decant the supernatant.
5. Resuspend the cell pellet in 500μL of PBS(F).
6. Spin down the resuspension at 3000rpm for 3 minutes.
7. Decant the supernatant.
8. Resuspend the cell pellet in another 500uL of PBS(F).*
9. Prepare the C6 Accuri Flow Cytometer by running a backflush cycle and a dH2O cycle.
10. Load the sample onto the sip.
11. Run the sample with 100,000 cell count.
12. Repeat for all samples and make sure to change data cells otherwise the old data is erased.
13. Once finished, run a cleaning cycle with Accuri approved cleaning solution, then run a dH2O cycle.
*For special cases do not resuspend all samples, instead resuspend immediately before running the sample through the flow cytometer.
Fluorescence Activated Cell Sorting
Final Preparations
Sample Prep:
Spin down samples and negative control (5000 RPM, 1 min), keeping in mind the library size. Aspirate off supernatant. Resuspend in PBSF. Spin down cells. Aspirate off supernatant. Resuspend in PBSF.
1. Open the “iGEM Template” file in the FACS Software and change name to current date and sort cycle
2. Make sure the stream is stable (look for green light in bottom right corner). If not, run the Sort Calibration order.
3. Run Negative Control from the PyE1 cells.
a. Load cells onto carrier and into the machine. Press play button on screen.
b. Set gate around lower left quadrant of cells to ensure single cell analysis using Forward Scatter Area and Side Scatter Area as your axes. Make sure oval gate covers around 80% of cell population.
c. Set second gate on the first gated population by double-clicking on the gated population and using Forward Scatter Height and Forward Scatter Width as your axes. You will notice two distinct populations. Try to focus on the single cell portion of the plot.
NOTE: If you see a large portion of the second gated population existing near the upper right edge of the first gate, you may need to enlarge the first gate to fit more of the population.
d. Press record. Record 100,000 events and stop run. Move to Next Tube.
4. Run first control (Gene clone)
a. Follow step 3 to run the first control
5. Run first library sample
a. Follow steps 3b and 3c to set first two gates correctly.
b. Set final gate for sort which includes top 1% of GFP producers from second gated population.
c. Use final gate to set up the sort.
d. Select sort conditions at the bottom of the screen.
e. Insert and load collection tube.
f. Record 100,000 events, and sort 10x the library size
6. Run Bleach and diH2O through FACS to avoid cross-contamination.
Protein Expression
Overnight Cultures
1. Add 25mL TB and 25uL Kan to a 250mL baffled flask
2. Stab a glycerol stock with a p1000 pipette and swirl in the flask of media
3. Put flask in 37C shaker overnight
Protein Expression
1. Add 500μL 1000x Kanamycin to 500mL TB in 2L baffled flask
2. Transfer 10mL overnight culture to TB
3. Shake at 37C (DO WE NEED THE RPM?)
4. Remove flask from shaker when optical density is between 0.5 and 0.8
5. Allow flask to rest at room temp for 30 min
6. Add 125μL 1M IPTG
7. Shake flask at 18C overnight
Protein Extraction and Purification
1. Transfer cell culture to centrifuge flask
2. Centrifuge culture at 4000g for 10 min
3. Discard supernatant
4. Resuspend pellet in 25mL lysis buffer
5. Add 250μL of 100x PMSF, 250μL of 100mg/mL lysozyme, and 250μL of 10mg/mL DNAse
6. Sonicate sample with 0.25inch probe for 5 min at 70% amplitude with 20 sec on and off pulses
7. Take 50μL total sample
8. Transfer lysate to SS-34 centrifuge tube
9. Centrifuge for 30 min at 18000g
10. Take 50μL soluble sample
Nickel Nitrotriacetic Acid Chromatography (Nickel-NTA Chromatography)
1. Add 5mL 50%(v/v) nickel resin in ethanol to 25mL column and allow to settle to ~2.5mL(CV)
2. Rinse with 10CV H20
3. Equilibrate with 10CV lysis buffer
4. Load sample onto column
5. Wash column with 15CV lysis buffer
6. Perform 2 additional wash steps with 15CV
7. Elute sample in 10CV elution buffer and collect eluate
8. Take 50μL pure sample
Size Exclusion Chromatography (S.E.C.)
1. Concentrate sample to as high as possible without inducing protein aggregation
2. Pre-equilibrate Superdex 75 column with 48 ml PBS
3. Inject 500 ul sample onto column
4. Run 36 ml PBS through column at 0.5 ml/min, collecting 1 ml fractions
5. Verify presence of protein in fractions by measuring concentration on Nanodrop and running SDS-PAGE (15 kDa protein should elute at ~13 ml)
6. Pool fractions containing protein
Stability Analysis
Circular Dichroism: Wavelength Scan
1. Load 1mm cuvette with 400uL protein solution onto CD
2. Take wavelength scan
260nm-190nm
sample every 1nm
averaging time 3sec
1 scan
step scan
25C
3. Record wavelength which gives strongest signal (222nm)
Circular Dichroism: Guanidine Melt
1. Load 1cm cuvette containing 1.996mL of 0.05mg/mL protein solution and stirrer onto CD
2. Prepare 8mL of 0.05mg/mL protein in concentrated guanidine solution
3. Set up Automixer with guanidine solution on one syringe and waste tube on other syringe
4. Titrate up to 6 M guanidine, taking a CD measurement at 222 nm every 0.15 M interval
5. Also measure the fluorescence at 280 nm to ensure the total protein concentration is not changing
