Team:ETH Zurich/InterLab

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This year the fifth international iGEM InterLab study took place. The goal of this years InterLab study is to establish a reliable framework to compare fluerecence measurements from different laboratorys. To achieve this goal it is essential to identify and correct sources of systematic variability in synthetic biology measurements. As previous InterLab studies have shown, it is possible to dramatically reduce systematic variability in GFP fluerescence measurements among different laboratory if the intensity is measured in absolute fluorescence units and calibrated against a known concentration of fluorescent molecule. However, when it comes to bulk measurments of cells expressing a fluerescent molecule the number of cells in the sample remains a source of considerable systematic variability. This years InterLab study aims to overcome this limitation by determining a mean expression level of GFP per cell, rather than an absolute fluorescence measurement.

Two orthogonal approaches are applied to determine the number of cells in a sample:

  1. Converting between absorbance of cells to absorbance of a known concentration of beads.
  2. Counting colony-forming units (CFUs) from the sample.

The cell count is later used to normalize the measured absolute fluerescence.
In the following sections we present the most important data from the InterLab study and the used protocoles. For further information please refere to the official iGEM InterLab study webpage.

General Information
Plasmides
The plasmides used for the InterLab study are specified in Table 1.
Test Devices for the InterLab study
Device Part Number Plate
Negative control BBa_R0040 Kit Plate 7 Well 2D
Positive control BBa_I20270 Kit Plate 7 Well 2B
Test Device 1 BBa_J364000 Kit Plate 7 Well 2F
Test Device 2 BBa_J364001 Kit Plate 7 Well 2H
Test Device 3 BBa_J364002 Kit Plate 7 Well 2J
Test Device 4 BBa_J364007 Kit Plate 7 Well 2L
Test Device 5 BBa_J364008 Kit Plate 7 Well 2N
Test Device 6 BBa_J364009 Kit Plate 7 Well 2P
Measurement Device
The InterLab Study was performed with a Tecan Infinite Nano. Refere to Table 2 for further information.
Measurement Device Specification
Feature Specification
Instrument brand and model Tecan Infinite Nano
Read mode Fluorescence/Absorbance
Pathlength correction Yes, disabled
Temperature 24°C
Filters excitation 485 nm (20 nm), emission 535 nm (25 nm)
Optics Top
Calibration
OD600 Reference point - LUDOX Protocol
LUDOX CL-X (45% colloidal silica suspension) was used as single point reference to obtain a conversion factor to transform the OD600 from our device into a comparable measurment. In the table below we present the results from these measurements.
OD600 Reference Point
LUDOX CL-X H2O
Replicate 1 0.063 0.034
Replicate 2 0.055 0.035
Replicate 3 0.060 0.034
Replicate 4 0.056 0.035
Arith. Mean 0.059 0.035
Corrected Abs600 0.024  
Reference OD600 0.063  
OD600/Abs600 2.630
Particle Standard Curve - Microsphere Protocol
Particel Standard Curve
Number of Particles 2.35E+08 1.18E+08 5.88E+07 2.94E+07 1.47E+07 7.35E+06 3.68E+06 1.84E+06 9.19E+05 4.60E+05 2.30E+05 0
Replicate 1 0.878 0.499 0.277 0.159 0.097 0.066 0.051 0.042 0.038 0.036 0.035 0.034
Replicate 2 0.879 0.502 0.280 0.159 0.097 0.066 0.050 0.042 0.039 0.036 0.036 0.034
Replicate 3 0.874 0.500 0.281 0.160 0.097 0.066 0.050 0.043 0.038 0.037 0.035 0.034
Replicate 4 0.876 0.500 0.281 0.160 0.097 0.066 0.050 0.042 0.038 0.036 0.035 0.034
Arith. Mean 0.877 0.500 0.280 0.159 0.097 0.066 0.050 0.042 0.038 0.036 0.035 0.034
Arith. Std.Dev. 0.002 0.001 0.002 0.000 0.000 0.000 0.001 0.001 0.000 0.000 0.000 0.000
Arith. Net Mean 0.843 0.466 0.245 0.125 0.063 0.032 0.016 0.008 0.004 0.002 0.001
Particles / OD
Number of Particles 2.35E+08 1.18E+08 5.88E+07 2.94E+07 1.47E+07 7.35E+06 3.68E+06 1.84E+06 9.19E+05 4.60E+05 2.30E+05
Mean particles / Abs600 2.79E+08 2.52E+08 2.40E+08 2.35E+08 2.35E+08 2.33E+08 2.30E+08 2.28E+08 2.28E+08 2.27E+08 2.24E+08
Mean of med-high levels: 2.39E+08                    
Background
Particle Standard Curve Log Scale
Background
Particle Standard Curve Linear Scale
Particle Standard Curves
Fluorescence standard curve - Fluorescein Protocol
Fluorescein Standard Curve
Fluorescein uM 10.00 5 2.5 1.25 0.625 0.313 0.156 0.078 0.039 0.0195 0.0098 0
Replicate 1 3.512E+04 2.443E+04 1.439E+04 7.764E+03 4.048E+03 2.037E+03 1.044E+03 5.210E+02 2.620E+02 1.330E+02 6.800E+01 1.000E+00
Replicate 2 3.327E+04 2.487E+04 1.430E+04 7.829E+03 4.087E+03 2.013E+03 1.016E+03 4.980E+02 2.530E+02 1.260E+02 6.300E+01 1.000E+00
Replicate 3 4.086E+04 2.489E+04 1.452E+04 7.891E+03 4.087E+03 2.059E+03 1.038E+03 5.100E+02 2.600E+02 1.310E+02 6.600E+01 1.000E+00
Replicate 4 3.813E+04 2.281E+04 1.503E+04 8.506E+03 4.626E+03 2.436E+03 1.267E+03 6.560E+02 3.470E+02 1.820E+02 9.600E+01 1.000E+00
Arith. Mean 3.684E+04 2.425E+04 1.456E+04 7.998E+03 4.212E+03 2.136E+03 1.091E+03 5.463E+02 2.805E+02 1.430E+02 7.325E+01 1.000E+00
Arith. Std.Dev. 3.345E+03 9.813E+02 3.289E+02 3.429E+02 2.766E+02 2.007E+02 1.178E+02 7.377E+01 4.450E+01 2.617E+01 1.531E+01 0.000E+00
Arith. Net Mean 3.684E+04 2.425E+04 1.456E+04 7.997E+03 4.211E+03 2.135E+03 1.090E+03 5.453E+02 2.795E+02 1.420E+02 7.225E+01
Fluorescein/a.u.
Fluorescein uM 10.00 5.00 2.50 1.25 0.63 0.31 0.16 0.08 0.04 0.02 0.01
uM Fluorescein/a.u. 2.71E-04 2.06E-04 1.72E-04 1.56E-04 1.48E-04 1.46E-04 1.43E-04 1.43E-04 1.40E-04 1.38E-04 1.35E-04
Mean uM fluorescein / a.u.:   1.66E-04                  
MEFL / a.u.:   9.99E+08                
Background
Fluorescein Standard Curve Log Scale
Background
Fluorescein Standard Curve Logarithmic Scale
Fluorescein Standard Curves
Cell measurement
Colony Forming Units
Fluorescence Raw Readings at Hour 0
Hour 0: Neg. Control Pos. Control Device 1 Device 2 Device 3 Device 4 Device 5 Device 6 LB + Chlor (blank)
Colony 1, Replicate 1 27 53 264 111 23 251 168 62 22
Colony 1, Replicate 2 19 53 269 93 23 260 184 59 23
Colony 1, Replicate 3 22 52 273 96 24 255 185 66 23
Colony 1, Replicate 4 22 55 265 93 24 262 200 61 23
Colony 2, Replicate 1 34 51 283 89 23 259 120 57 24
Colony 2, Replicate 2 23 58 257 87 23 268 130 60 22
Colony 2, Replicate 3 22 60 271 93 23 270 129 60 22
Colony 2, Replicate 4 25 58 289 96 25 282 135 61 23
Abs600 Raw Readings at Hour 0
Hour 0: Neg. Control Pos. Control Device 1 Device 2 Device 3 Device 4 Device 5 Device 6 LB + Chlor (blank)
Colony 1, Replicate 1 0.0639 0.0588 0.0582 0.0601 0.0575 0.0589 0.0552 0.0586 0.0397
Colony 1, Replicate 2 0.0562 0.0586 0.0630 0.0571 0.0612 0.0603 0.0567 0.0585 0.0395
Colony 1, Replicate 3 0.0575 0.0580 0.0590 0.0582 0.0583 0.0589 0.0557 0.0594 0.0397
Colony 1, Replicate 4 0.0584 0.0594 0.0583 0.0581 0.0583 0.0593 0.0568 0.0585 0.0395
Colony 2, Replicate 1 0.0647 0.0578 0.0592 0.0576 0.0572 0.0588 0.0486 0.0578 0.0426
Colony 2, Replicate 2 0.0580 0.0594 0.0565 0.0573 0.0571 0.0587 0.0503 0.0592 0.0412
Colony 2, Replicate 3 0.0571 0.0603 0.0580 0.0581 0.0573 0.0595 0.0500 0.0592 0.0396
Colony 2, Replicate 4 0.0588 0.0596 0.0580 0.0590 0.0592 0.0613 0.0507 0.0589 0.0394
Fluorescence Raw Readings at Hour 6
Hour 6: Neg. Control Pos. Control Device 1 Device 2 Device 3 Device 4 Device 5 Device 6 LB + Chlor (blank)
Colony 1, Replicate 1 30 999 516 1914 74 1479 460 952 22
Colony 1, Replicate 2 32 1090 546 2064 79 1457 514 1043 23
Colony 1, Replicate 3 32 1089 568 2099 79 1491 506 1054 23
Colony 1, Replicate 4 32 1089 546 2105 80 1526 512 1047 22
Colony 2, Replicate 1 34 1112 570 1817 57 1701 407 1123 24
Colony 2, Replicate 2 35 1085 601 1767 60 1656 391 1127 23
Colony 2, Replicate 3 35 1126 620 1783 59 1662 406 1123 23
Colony 2, Replicate 4 34 1098 596 1791 59 1721 403 1144 24
Abs600 Raw Readings at Hour 6
Hour 6: Neg. Control Pos. Control Device 1 Device 2 Device 3 Device 4 Device 5 Device 6 LB + Chlor (blank)
Colony 1, Replicate 1 0.4669 0.4309 0.0698 0.4685 0.5371 0.2058 0.0742 0.4678 0.0445
Colony 1, Replicate 2 0.4925 0.4496 0.0711 0.4913 0.5618 0.2010 0.0755 0.5018 0.0413
Colony 1, Replicate 3 0.4904 0.4542 0.0709 0.5042 0.5552 0.2048 0.0774 0.5099 0.0400
Colony 1, Replicate 4 0.4932 0.4546 0.0727 0.5049 0.5525 0.2078 0.0789 0.5126 0.0396
Colony 2, Replicate 1 0.4932 0.4858 0.0784 0.4988 0.4786 0.2288 0.0699 0.5129 0.0393
Colony 2, Replicate 2 0.5286 0.4903 0.0745 0.4955 0.5086 0.2375 0.0698 0.5161 0.0397
Colony 2, Replicate 3 0.5068 0.4970 0.0756 0.4994 0.5045 0.2363 0.0710 0.5080 0.0396
Colony 2, Replicate 4 0.4992 0.4837 0.0800 0.5047 0.5059 0.2411 0.0706 0.5311 0.0396
Protocols
Competent Cell Production (from stock)
Introduction
Detergent is a major inhibitor of competent cell growth and transformation. Glass and plastic must be detergent free for these protocols. The easiest way to do this is to avoid washing glassware with detergent, and simply rinse it out. Autoclaving glassware filled 3/4 with DI water is an effective way to remove most detergent residue. Media and buffers should be prepared in detergent free glassware and cultures grown up in detergent free glassware.
Materials
  • CCMB80 buffer
  • SOB
  • Detergent-free, sterile glassware and plasticware
  • Table-top OD600nm spectrophotometer
Procedure
  1. Prechill 250mL centrifuge tubes and screw cap tubes before use.

  2. Ethanol treat all working areas for sterility.

  3. Inoculate 250 ml of SOB medium with 1 ml vial of seed stock and grow at 20°C to an OD600nm of 0.3. Use the "cell culture" function on the Nanodrop to determine OD value. OD value = 600nm Abs reading x 10.
    This takes approximately 16 hours. Controlling the temperature makes this a more reproducible process, but is not essential. Room temperature will work. You can adjust this temperature somewhat to fit your schedule Aim for lower, not higher OD if you can't hit this mark
  4. Fill an ice bucket halfway with ice. Use the ice to pre-chill as many flat bottom centrifuge bottles as needed.

  5. Transfer the culture to the flat bottom centrifuge tubes. Weigh and balance the tubes using a scale
. Try to get the weights as close as possible, within 1 gram.
  6. Centrifuge at 3000g at 4°C for 10 minutes in a flat bottom centrifuge bottle.
 Flat bottom centrifuge tubes make the fragile cells much easier to resuspend
  7. Decant supernatant into waste receptacle, bleach before pouring down the drain.
  8. Gently resuspend in 80 ml of ice cold CCMB80 buffer

    Pro tip: add 40ml first to resuspend the cells. When cells are in suspension, add another 40ml CCMB80 buffer for a total of 80ml Pipet buffer against the wall of the centrifuge bottle to resuspend cells. Do not pipet directly into cell pellet! After pipetting, there will still be some residual cells stuck to the bottom. Swirl the bottles gently to resuspend these remaining cells
  9. Incubate on ice for 20 minutes

  10. Centrifuge again at 3000G at 4°C. Decant supernatant into waste receptacle, and bleach before pouring down the drain.

  11. Resuspend cell pellet in 10 ml of ice cold CCMB80 buffer.
If using multiple flat bottom centrifuge bottles, combine the cells post-resuspension
  12. Use Nanodrop to measure OD of a mixture of 200 μl SOC and 50 μl of the resuspended cells
. Use a mixture of 200 μl SOC and 50 μl CCMB80 buffer as the blank
  13. Add chilled CCMB80 to yield a final OD of 1.0-1.5 in this test.
  14. Incubate on ice for 20 minutes. Prepare for aliquoting
.
    Make labels for aliquots. Use these to label storage microcentrifuge tubes/microtiter plates Prepare dry ice in a separate ice bucket. Pre-chill tubes/plates on dry ice.
  15. Aliquot into chilled 2ml microcentrifuge tubes or 50 μl into chilled microtiter plates
  16. Store at -80°C indefinitely.Flash freezing does not appear to be necessary
  17. Perform test transformations to calculate your competent cell efficiency
.
    Thawing and refreezing partially used cell aliquots dramatically reduces transformation efficiency by about 3x the first time, and about 6x total after several freeze/thaw cycles. Good cells should yield around 100 - 400 colonies Transformation efficiency is (dilution factor=15) x colony count x 105/µgDNA We expect that the transformation efficiency should be between 1.5x108 and 6x108cfu/µgDNA
Evaluation
Bla Bla
Results
Did not work at all!
Competent Cell Test Kit
Introduction
Before using your competent cells in an experiment, use the Competent Cell Test Kit to test the efficiency of your competent cells! The kit includes two vials of dried-down purified plasmid DNA from BBa_J04450 (RFP construct) in plasmid backbone pSB1C3. When resuspended with 50uL dH2O, the vials will result in different concentrations: 100 pg/µL, 10 pg/µL. Perform transformations with each of these to determine how efficient your competent cells are. It is important to have efficient competent cells because transformations performed with ligation products usually do not yield as many colonies due to the low DNA concentration in the ligation mixture. This means that you may see different results doing this test than you will at the end of the 3A Assembly protocol, or any other ligation. You can use the following suggested protocol for heat-shock transformation to test your chemically-competent E. coli cells.
Materials
  • 70% ethanol
  • Paper towels
  • Lab marker / Sharpie
  • 1.5 mL microcentrifuge tubes
  • Container for ice
  • Ice
  • Competent cell aliquot(s)
  • Competent Cell Test Kit
  • Agar plates with chloramphenicol
  • 42°C Waterbath (or hot water source and thermometer)
  • 37°C Incubators (oven and shaker)
  • SOC media
  • Sterile glass beads or sterile cell spreader
  • Pipettor
  • Pipette tips
Procedure
  1. Clean your working area by wiping down with 70% ethanol.

  2. Thaw competent cells on ice. Label one 1.5 mL microcentrifuge tubes for each transformation and then pre-chill by plcaacing the tubes on ice. Do triplicates (3 each) of each concentration if possible, so you can calculate an average colony yield.
  3. Spin down the DNA tubes from the Competent Cell Test Kit/Transformation Efficiency Kit to collect all of the DNA into the bottom of each tube prior to use. A quick spin of 20-30 seconds at 8,000-10,000 rpm will be sufficient. Note: You should resuspend the DNA in each tube with 50 µL dH2O.
  4. Pipet 1 µL of DNA into each microcentrifuge tube.
  5. Pipet 50 µL of competent cells into each tube. Flick the tube gently with your finger to mix.
  6. Incubate on ice for 30 minutes. Pre-heat waterbath now to 42°C. Otherwise, hot water and an accurate thermometer works, too!
  7. Heat-shock the cells by placing into the waterbath for 45 seconds (no longer than 1 min). Be careful to keep the lids of the tubes above the water level, and keep the ice close by.
  8. Immediately transfer the tubes back to ice, and incubate on ice for 5 minutes.
  9. Add 950 µL of SOC media per tube, and incubate at 37°C for 1 hour shaking at 200-300rpm. Prepare the agar plates during this time: label them, and add sterile glass beads if using beads to spread the mixture.
  10. Pipet 100 µL from each tube onto the appropriate plate, and spread the mixture evenly across the plate. Incubate at 37°C overnight or approximately 16 hours. Position the plates with the agar side at the top, and the lid at the bottom.
  11. Count the number of colonies on a light field or a dark background, such as a lab bench. Use the following equation to calculate your competent cell efficiency. If you've done triplicates of each sample, use the average cell colony count in the calculation. Efficiency (in cfu/µg) = [colonies on plate (cfu) / Amount of DNA plated (ng)] x 1000 (ng/µg). Note: The measurement "Amount of DNA plated" refers to how much DNA was plated onto each agar plate, not the total amount of DNA used per transformation. You can calculate this number using the following equation: Amount of DNA plated (ng) = Volume DNA added (1 µL) x concentration of DNA (refer to vial, convert to ng/µL) x [volume plated (100 µL) / total reaction volume (1000 µL)]
Results
Competent cells should have an efficiency of 1.5x10^8 to 6x10^8 cfu/µg DNA, where "cfu" means "colony-forming unit" and is a measurement of cells.
LUDOX Protocol
Introduction
Protocol provided by the iGEM fundation to perform the LUDOX calibration
Materials
  • 1ml LUDOX CL-X (provided in kit)
  • ddH20 (provided by team)
  • 96 well plate, black with clear flat bottom preferred (provided by team)
Procedure
Note that the the Ludox tube was stored at Room temperature, not at +4 degrees. (not in the fridge). This has been communicated to the iGEM foundation.
  1. Add 100 μl LUDOX into wells A1, B1, C1, D1
  2. Add 100 μl of dd H2O into wells A2, B2, C2, D2
  3. Measure absorbance at 600 nm of all samples in the measurement mode you plan to use for
  4. cell measurements
  5. Record the data in the table below or in your notebook
  6. Import data into Excel sheet provided (OD600 reference point tab​)
Results
See Table 1
Microsphere Protocol
Introduction
Protocol provided by the iGEM fundation to perform the Microsphere calibration curve.
Materials
  • 300 μL Silica beads - Microsphere suspension (provided in kit, 4.7 x 10^8 microspheres)
  • ddH20 (provided by team)
  • 96 well plate, black with clear flat bottom preferred (provided by team)
Procedure
  1. Obtain the tube labeled “Silica Beads” from the InterLab test kit and vortex vigorously for 30 seconds. NOTE: Microspheres should NOT be stored at 0°C or below​, as freezing affects the properties of the microspheres. If you believe your microspheres may have been frozen, please contact the iGEM Measurement Committee for a replacement (measurement at igem dot org).
  2. Immediately pipet 96 μL microspheres into a 1.5 mL eppendorf tube.
  3. Add 904 μL of ddH2O to the microspheres
  4. Vortex well. This is your Microsphere Stock Solution.
  5. Add 100 μl of ddH​2​O​ into wells A2, B2, C2, D2....A12, B12, C12, D12
  6. Vortex the tube containing the stock solution of microspheres vigorously for 10 seconds
  7. Immediately add 200 μl​ of microspheres stock​ solution into A1
  8. Transfer 100 μl of microsphere stock solution from A1 into A2.
  9. Mix A2 by pipetting up and down 3x and transfer 100 μl into A3…
  10. Mix A3 by pipetting up and down 3x and transfer 100 μl into A4...
  11. Mix A4 by pipetting up and down 3x and transfer 100 μl into A5...
  12. Mix A5 by pipetting up and down 3x and transfer 100 μl into A6...
  13. Mix A6 by pipetting up and down 3x and transfer 100 μl into A7...
  14. Mix A7 by pipetting up and down 3x and transfer 100 μl into A8...
  15. Mix A8 by pipetting up and down 3x and transfer 100 μl into A9...
  16. Mix A9 by pipetting up and down 3x and transfer 100 μl into A10...
  17. Mix A10 by pipetting up and down 3x and transfer 100 μl into A11...
  18. Mix A11 by pipetting up and down 3x and transfer 100 μl into liquid waste
  19. Repeat dilution series for rows B, C, D
  20. IMPORTANT! ​Re-Mix (Pipette up and down) each row of your plate immediately
  21. before putting in the plate reader! (This is important because the beads begin to settle to
  22. the bottom of the wells within about 10 minutes, which will affect the measurements.) Take
  23. care to mix gently and avoid creating bubbles on the surface of the liquid.
  24. Measure Abs600 of all samples in instrument
  25. Record the data in your notebook
  26. Import data into Excel sheet provided (particle standard curve tab​)
Results
See Table 2 and Table 3
Fluorescein Protocol
Introduction
Protocol provided by the iGEM fundation to perform the Microsphere calibration curve.
Materials
  • Fluorescein (provided in kit)
  • 10ml 1xPBS pH 7.4-7.6 (phosphate buffered saline; provided by team)
  • 96 well plate, black with clear flat bottom (provided by team)
Procedure
  1. Spin down fluorescein kit tube to make sure pellet is at the bottom of tube.
  2. Prepare 10x fluorescein stock solution (100 μM) by resuspending fluorescein in 1mL of 1xPBS. [ Note​: it is important that the fluorescein is properly dissolved. To check this, after the resuspension you should pipette up and down and examine the solution in the pipette tip – if any particulates are visible in the pipette tip continue to mix the solution until they disappear.]
  3. Dilute the 10x fluorescein stock solution with 1xPBS to make a 1x fluorescein solution with concentration 10 μM: 100 μL of 10x fluorescein stock into 900 μL 1xPBS
  4. Add 100 μl of PBS​ into wells A2, B2, C2, D2....A12, B12, C12, D12
  5. Add 200 μl​ of fluorescein 1x stock​ solution into A1, B1, C1, D1
  6. Transfer 100 μl of fluorescein stock solution from A1 into A2.
  7. Mix A2 by pipetting up and down 3x and transfer 100 μl into A3…
  8. Mix A3 by pipetting up and down 3x and transfer 100 μl into A4...
  9. Mix A4 by pipetting up and down 3x and transfer 100 μl into A5...
  10. Mix A5 by pipetting up and down 3x and transfer 100 μl into A6...
  11. Mix A6 by pipetting up and down 3x and transfer 100 μl into A7...
  12. Mix A7 by pipetting up and down 3x and transfer 100 μl into A8...
  13. Mix A8 by pipetting up and down 3x and transfer 100 μl into A9...
  14. Mix A9 by pipetting up and down 3x and transfer 100 μl into A10...
  15. Mix A10 by pipetting up and down 3x and transfer 100 μl into A11...
  16. Mix A11 by pipetting up and down 3x and transfer 100 μl into liquid waste
  17. Repeat dilution series for rows B, C, D
  18. Measure fluorescence of all samples in instrument
  19. Record the data in your notebook
  20. Import data into Excel sheet provided (fluorescein standard curve tab​)
Results
See Table 4 and Table 5
Cell measurement protocol
Introduction
Protocol provided by the iGEM fundation to perform the cell measurement.
Materials
  • Competent cells (Escherichia coli strain DH5α)
  • LB (Luria Bertani) media
  • Chloramphenicol (stock concentration 25 mg/mL dissolved in EtOH)
  • 50 ml Falcon tube (or equivalent, preferably amber or covered in foil to block light)
  • Incubator at 37°C
  • 1.5 ml eppendorf tubes for sample storage
  • Ice bucket with ice
  • Micropipettes and tips
  • 96 well plate, black with clear flat bottom preferred (provided by team)
Procedure
  1. Day 1​: transform Escherichia coli DH5α with the plasmides specified above
  2. Day 2​: Pick 2 colonies from each of the transformation plates and inoculate in 5-10 mL LB medium +Chloramphenicol. Grow the cells overnight (16-18 hours) at 37°C and 220 rpm.
  3. Day 3​: Cell growth, sampling, and assay
  4. Make a 1:10 dilution of each overnight culture in LB+Chloramphenicol (0.5mL of
  5. culture into 4.5mL of LB+Chlor)
  6. Measure Abs600 of these 1:10 diluted cultures
  7. Record the data in your notebook
  8. Dilute the cultures further to a target Abs600 of 0.02 in a final volume of 12 ml LB medium + Chloramphenicol in 50 mL falcon tube (amber, or covered with foil to block light).
  9. Take 500 µL samples of the diluted cultures at 0 hours into 1.5 ml eppendorf tubes,prior to incubation. (At each time point 0 hours and 6 hours, you will take a sample from each of the 8 devices, two colonies per device, for a total of 16 eppendorf tubeswith 500 µL samples per time point, 32 samples total).
  10. Place the samples on ice.
  11. Incubate the remainder of the cultures at 37°C and 220 rpm for 6 hours.
  12. Take 500 µL samples of the cultures at 6 hours of incubation into 1.5 ml eppendorf tubes. Place samples on ice.
  13. At the end of sampling point you need to measure your samples (Abs600 and fluorescence measurement), see the below for details.
  14. Record data in your notebook
  15. Import data into Excel sheet provided (fluorescence measurement tab​)
Results
See Tables and Figures in cell measurement section
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