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Team:Washington/Protocols
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14. Quickly aliquot the cells into 1.7 mL cryogenic vials or 1.5 mL centrifuge tubes.** <br> | 14. Quickly aliquot the cells into 1.7 mL cryogenic vials or 1.5 mL centrifuge tubes.** <br> | ||
15. Store the competent cell aliquots at -80 °C. <br> | 15. Store the competent cell aliquots at -80 °C. <br> | ||
| - | + | <br> | |
*Streak in such a way that there should be individual colony growth and no clumps after the incubation. <br> | *Streak in such a way that there should be individual colony growth and no clumps after the incubation. <br> | ||
**We did this in a -20 °C cold room and using an automated repeater pipette. The volume of each aliquot depends on the number of transformations you intend to do at a time. <br> | **We did this in a -20 °C cold room and using an automated repeater pipette. The volume of each aliquot depends on the number of transformations you intend to do at a time. <br> | ||
Revision as of 22:48, 17 October 2014
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Contents |
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 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 in a 1 L bottle
Phosphate Buffered Saline (PBS) Solution
Mix the following in 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 in a 1 L bottle
PBSF (PBS for Flow)
Mix the following in a 1 L beaker:
- 25 mL 20X PBS, pH 7.4
- 475 mL H2O
- 2.5 g BSA (0.5%)*
Sterile filter in a 1L bottle and store at 4 °C
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
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
Add the following to a 500mL beaker and mix:
- 203 g guanidinium hydrogen chloride
- 250 mL PBS solution*
- Add dilute HCl to pH 7.4
*It is not necessary to filter or autoclave.
*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
Competent cells take two days to culture and aliquot.
Day 1:
1. Streak an aliquot of competent cells onto two LB-plates without antibiotics.*
2. Incubate at 37 °C overnight.
Day 2:
1. In two 250 mL baffle flasks add 50 mL of SOB media.
2. Scrape as many single colonies into either flask.
3. Incubate and shake at 37 °C and 250 rpm for 2-3 hours.
4. Check the optical density of the cells at 600 nm after 2 hours.
5. Stop incubation when cultures reach approximately 0.5 optical density.
6. Add the contents of the flask into separate 50 mL flat bottomed centrifuge tubes.
7. Spin down the cells at 2500 rpm at 4 °C for 15 minutes.
8. Decant the supernatant.
9. Resuspend the cells in 16 mL of CCMB by pipetting or gently vortexing.
10. Incubate the cells on ice for 20 minutes.
11. Spin down the cells at 2500 rpm at 4 °C for 10 minutes.
12. Decant the supernatant.
13. Resuspend the cells in 4 mL of CCMB.
14. Quickly aliquot the cells into 1.7 mL cryogenic vials or 1.5 mL centrifuge tubes.**
15. Store the competent cell aliquots at -80 °C.
*Streak in such a way that there should be individual colony growth and no clumps after the incubation.
**We did this in a -20 °C cold room and using an automated repeater pipette. The volume of each aliquot depends on the number of transformations you intend to do at a time.
Note: After removing the cells from incubation keep them on ice or as cold as possible.
Chemically Competent Cell Transformations
1. Thaw competent E. coli cells on ice (XL1-Blue or XL10-Gold).*
2. Add 50 μL of competent cells to sterile 14 mL culture tube.
3. Add 1 μL (~100-200 ng)* of the mini-prep to each culture tube.
4. Equilibrate the cells on ice for 10 minutes.
5. Heat shock the cells at 42 °C for 30-45 seconds.**
6. Immediately place the cells back on ice for 3 minutes.
7. Add 250 μL LB media without antibiotics and shake at 250 rpm and 37 °C for 30 minutes.
8. Spread 10 μL and 290 μL on an appropriate LB-antibiotic plate.
9. Invert the plate and incubate at 37 °C 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 14 mL round-bottom tube, add 3 mL of LB and 3 μL of 1000X antibiotic(s).
2. Pick one isolated colony, 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 37 °C at 250 rpm for 12-16 hours and no longer than 20 hours.
DNA Extraction and Mini-Preps
All DNA Mini-Preps were prepared using EPOCH Mini-Prep Kits and following the supplied protocols.
Glycerol Stocks
1. Take 1-2 mL from an overnight culture and transfer into a 1.5 mL centrifuge tube.
2. Spin down the culture at 3000 rpm for 3 minutes.
3. Decant the supernatant.
4. Resuspend the cells in 500 μL of 40% glycerol and 500 μL of LB (no antibiotics) or water.
5. Transfer the resuspension to a cryogenic vial.
6. Store the glycerol stock at -80 °C.
Saccharomyces cerevisiae (PyE1 Yeast)
Chemically Competent Cell Cultures
This process take four days in lab with a one day wait for incubation.
Day 1:
1. Streak yeast cells onto a YPD plate.*
2. Invert the plate and incubate at 30 °C for 2 days.
Day 3:
1. Add 50 mL of YPD liquid media into a 250 mL baffle flask.
2. Swipe as many individual colonies as you can see into the YPD media.**
3. Incubate and shake the culture at 30 °C at 250 rpm overnight approximately 24 hours.
Day 4:
1. Take an optical density measurement.
2. In three 250 mL 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 30 °C at 250 rpm until the optical density reaches 1.2-1.6.
5. Collect each culture into separate 50 mL flat-bottomed centrifuge tubes.
6. Spin down the cells at 4000 x g for 5 minutes at 4 °C.
7. Decant the supernatant.
8. Resuspend the cells in 100 mL total for all three culture of dH2O.
9. Combine the suspensions into two 50 mL flat-bottomed centrifuge tubes.
10. Spin down the cells as above.
11. Decant the supernatant.
12. Resuspend each in 3 mL of 100 mM lithium acetate.
13. Transfer both cultures into a single 15 mL conical centrifuge tube.
14. Spin down the cells at 3000 rpm for 5 minutes.
15. Resuspend the cells in 0.75 mL of 100 mM lithium acetate, total volume is roughly 2 mL.
16. Qualitatively bring up the volume to 3.5 mL by adding 40% glycerol.
17. Aliquot the cells into 1.5 mL centrifuge tubes or 1.7 mL 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 50 μL aliquot of yeast competent cells were made. Furthermore, this protocol prepares enough cells for six yeast transformations.
1. Add the following to 50 μL of yeast competent cells:
240 μL of polyethylene glycol - 3350 (PEG-3350)
36 μL of 1 M lithium acetate
32 μL of milliQ H2O
2. Mix the mixture by gently pipetting or vortexing.
3. Aliquot 59 μL of the mixture into a 0.2 mL microcentrifuge tube.
4. Add 1 μL (~100-200 ng) of DNA.*
5. Mix the mixture by gentle pipetting or vortexing.
6. Incubate the mixture at 30 °C for 30 minutes.
7. Heat shock the mixture at 42 °C for 20 minutes.
8. Spin down the cells in a microcentrifuge for ~1 minute.
9. Decant the supernatant.
10. Resuspend the cell pellets in 200 μL of dH2O.
11. Spin down the cells in a microcentrifuge for ~1 minute.
12. Resuspend the cell pellets in 200 μL of dH2O.
13. Plate 50-150 μL of the mixture onto an appropriate selective dropout media plate.
14. Invert and incubate at 30 °C for 2 days.
*The exact amount of DNA depends on the transformation efficiency of your competent cells.
Overnight Culturing
1. In a 14 mL round-bottomed culture tube add 1.8 mL selective dropout media and 0.2 mL 20% glucose.
2. Swipe 3 isolated yeast colonies and add them to the culture tube media.
3. Incubate and shake at 37 °C at 250 rpm for 2 days.
Note: You can also make 3 mL cultures (2.7 mL S.D. media and 0.3 mL 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 3 mL cultures (2.7 mL S.D. media and 0.3 mL 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 1-2 mL from an overnight culture and transfer into a 1.5 mL centrifuge tube.
2. Spin down the culture at 3000 rpm for 3 minutes.
3. Decant the supernatant.
4. Resuspend the cells in 500 μL of 40% glycerol and 500 μL of selective dropout media or water.
5. Transfer the resuspension to a cryogenic vial.
6. Store the glycerol stock at -80 °C.
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
