Team:CCU Taiwan/InterLab

INTERLAB

Background

   All of the 2018 iGEM teams are invited and encouraged to participate in the Fifth International InterLaboratory Measurement Study in synthetic biology.” Our team took part in this study which aimed to be one of the labs to make the calibration in different condition. In our case, we measured fluorescence using plate reader.


OD600 Reference Point - LUDOX Protocol

Materials：

  1. 1ml LUDOX CL-X (provided in kit)
  2. dd H2O (provided by team)
  3. 96 well plate, black with clear flat bottom preferred (provided by team)

Method：

  1. Add 100μL LUDOX into wells A1, B1, C1, D1
  2. Add 100μL 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 cell measurements
  4. Record the data in the table below or in your notebook
  5. Import data into Excel sheet provided ( OD600 reference point tab )

Data Result：

Figure1:LUDOX CL-X & H2O2





Particle Standard Curve - Microsphere Protocol

Materials：

  1. 300μL Silica beads - Microsphere suspension (provided in kit, 4.7 x 108 microspheres)
  2. dd H2O (provided by team)
  3. 96 well plate, black with clear flat bottom preferred (provided by team)

Method：

Prepare the Microsphere Stock Solution:

  1. Obtain the tube labeled “Silica Beads” from the Interlab test kit and vortex vigorously for 30 seconds
  2. Immediately pipet 96μL microspheres into a 1.5 mL eppendorf tube
  3. Add 904μL of dd H2O to the microspheres
  4. Vortex well

Data Result：

Figure2: Number of particles



Figure3: Number of particles



Figure4-1: Particle standard curve

Figure4-2: Particle standard curve(log scale)





Fluorescence Standard Curve - Fluorescein Protocol

Materials：

  1. Fluorescein (provided in kit)
  2. 10ml 1xPBS pH 7.4-7.6 (phosphate buffered saline; provided by team)
  3. 96 well plate, black with clear flat bottom (provided by team)

Method：

Prepare the fluorescein stock solution:

  1. Spin down fluorescein kit tube to make sure pellet is at the bottom of tube
  2. Prepare 10x fluorescein stock solution (100μM) by resuspending fluorescein in 1mL of 1xPBS
  3. Dilute the 10x fluorescein stock solution with 1xPBS to make a 1x fluorescein solution with concentration 10μM: 100μL of 10x fluorescein stock into 900μL 1xPBS

Prepare the serial dilutions of fluorescein:

  1. Add 100μL of PBS into wells A2, B2, C2, D2....A12, B12, C12, D12
  2. Add 200μL of fluorescein 1x stock solution into A1, B1, C1, D1
  3. Transfer 100μL of fluorescein stock solution from A1 into A2
  4. Mix A2 by pipetting up and down 3x and transfer 100μL into A3
  5. Mix A3 by pipetting up and down 3x and transfer 100μL into A4
  6. Mix A4 by pipetting up and down 3x and transfer 100μL into A5
  7. Mix A5 by pipetting up and down 3x and transfer 100μL into A6
  8. Mix A6 by pipetting up and down 3x and transfer 100μL into A7
  9. Mix A7 by pipetting up and down 3x and transfer 100μL into A8
  10. Mix A8 by pipetting up and down 3x and transfer 100μL into A9
  11. Mix A9 by pipetting up and down 3x and transfer 100μL into A10
  12. Mix A10 by pipetting up and down 3x and transfer 100μL into A11
  13. Mix A11 by pipetting up and down 3x and transfer 100μL into liquid waste
  14. Repeat dilution series for rows B, C, D
  15. Measure fluorescence of all samples in instrument
  16. Record the data in your notebook
  17. Import data into Excel sheet provided ( fluorescein standard curve tab )

Data Result：

Figure5: Fluorescein data results



Figure6-1: Fluorescein standard curve

Figure6-1:Fluorescein standard curve(log scale)



Figure7: Fluorescence arbitrary units





Cell Measurement Protocol

Materials：

  1. Competent cells (Escherichia coli strain DH5α)
  2. LB (Luria Bertani) media
  3. Chloramphenicol (stock concentration 25 mg/mL dissolved in EtOH)
  4. 50 ml Falcon tube (or equivalent, preferably amber or covered in foil to block light)
  5. Incubator at 37°C
  6. 1.5 ml eppendorf tubes for sample storage
  7. Ice bucket with ice
  8. Micropipettes and tips
  9. 96 well plate, black with clear flat bottom preferred (provided by team)

Method：

Day 1: transform Escherichia coli DH5α with these following plasmids (all in pSB1C3)

  1. Label 1.5ml tubes with part name or well location
  2. Take competent cells out of -80°C and thaw on ice (175µL commercial competent cell)
  3. Remove agar plates (containing the appropriate antibiotic) from storage at 4°C (without warm at 37℃ incubator)
  4. Pipette 25µL of commercial competent cell into 1.5ml tube
  5. Pipette 1µL of DNA into 1.5ml tube (without resuspended)
  6. Close 1.5ml tubes, incubate on ice for 35min ( should be 30min, but we take 5min to take 25µL competent cell for D3, and add it.)
  7. Heat shock tubes at 42°C for 45 sec
  8. Incubate on ice for 5min
  9. Pipette l LB media to 1000µL each transformation
  10. Incubate at 37°C for 1 hours, shaking at 200-300rpm
  11. Spin down cells at 6800g for 3mins and discard 800µL of the supernatant.Resuspend the cells in the remaining 200µL, and pipette eachtransformation onto petri plates


Day 2: Pick 2 colonies from each of the transformation plates and inoculate in 5-10mL LB medium with Chloramphenicol. Grow the cells overnight (16-18 hours) at 37°C and 220 rpm

Day 3: Cell growth, sampling, and assay

  1. Make a 1:10 dilution of each overnight culture in LB with Chloramphenicol (0.5mL of culture into 4.5mL of LB+Chlor)
  2. Measure Abs600 of these 1:10 diluted cultures
  3. Record the data in your notebook
  4. Dilute the cultures further to a target Abs600 of 0.02 in a final volume of 12 ml LB medium with Chloramphenicol in 50 mL falcon tube (amber, or covered with foil to block light)
  5. Take 500μL 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 tubes with 500μL samples per time point, 32 samples total)
    Place the samples on ice
  6. Incubate the remainder of the cultures at 37°C and 220 rpm for 6 hours
  7. Take 500μL samples of the cultures at 6 hours of incubation into 1.5 ml Eppendorf tubes.
    Place samples on ice
  8. At the end of sampling point you need to measure your samples (Abs600 and fluorescence measurement), see the below for details
  9. Record data in your notebook
  10. Import data into Excel sheet provided


Device Part Number Plate Location


Figure8: device part number plate location



Measurement

   Samples should be laid out according to the plate diagram below. Pipette 100μL of each sample into each well. From 500μL samples in a 1.5 ml eppendorf tube, 4 replicate samples of colony #1 should be pipetted into wells in rows A, B, C and D. Replicate samples of colony #2 should be pipetted into wells in rows E, F, G and H. Be sure to include 8 control wells containing 100μL each of only LB with chloramphenicol on each plate in column 9, as shown in the diagram below.
   Set the instrument settings as those that gave the best results in your calibration curves (no measurements off scale). If necessary you can test more than one of the previously calibrated settings to get the best data (no measurements off scale). Instrument temperature should be set to room temperature (approximately 20-25°C) if your instrument has variable temperature settings.



Data Results：

Fluorescence(a.u.) Raw Readings


Figure9: Fluorescence(a.u.) raw readings hour: 0



Figure10: Fluorescence(a.u.) raw readings hour: 6



Abs600 Raw Readings


Figure11: Abs600 raw readings hour: 0



Figure12: Abs600 raw readings hour: 6





Colony Forming Units per 0.1 OD600 E. coli cultures

Method：

Step 1: Starting Sample Preparation

  1. Measure the OD600 of your cell cultures, making sure to dilute to the linear detection range of your plate reader, e.g. to 0.05 – 0.5 OD600 range. Include blank media (LB + Cam) as well. For an overnight culture (16-18 hours of growth), we recommend diluting your culture 1:8 (8-fold dilution) in LB + Cam before measuring the OD600. Preparation: Add 25μL culture to 175μL LB + Cam in a well in a black 96-well plate, with a clear, at bottom. Recommended plate setup is below. Each well should have 200μL
  2. Dilute your overnight culture to OD600 = 0.1 in 1mL of LB + Cam media. Do this in triplicate for each culture.
  3. Check the OD600 and make sure it is 0.1 (minus the blank measurement).

Step 2: Dilution Series Instructions

Do the following serial dilutions for your triplicate Starting Samples you prepared in Step 1. You should have 12 total Starting Samples - 6 for your Positive Controls and 6 for your Negative Controls.

For each Starting Sample (total for all 12 showed in italics in paraenthesis)

  1. You will need 3 LB Agar + Cam plates (36 total).
  2. Prepare three 2.0 mL tubes (36 total) with 1900μL of LB + Cam media for Dilutions 1, 2, and 3 (see figure below).
  3. Prepare two 1.5 mL tubes (24 total) with 900μL of LB + Cam media for Dilutions 4 and 5 (see figure below)..
  4. Label each tube according to the figure below (Dilution 1, etc.) for each Starting Sample.
  5. Pipet 100μL of Starting Culture into Dilution 1. Discard tip. Do NOT pipette up and down. Vortex tube for 5-10 secs.
  6. Repeat Step 5 for each dilution through to Dilution 5 as shown below.
  7. Aseptically spead plate 100μL on LB + Cam plates for Dilutions 3, 4, and 5.
  8. Incubate at 37°C overnight and count colonies after 18-20 hours of growth.

Step 3: CFU/mL/OD Calculation Instructions

  1. Count the colonies on each plate with fewer than 300 colonies.
  2. Multiple the colony count by the Final Dilution Factor on each plate.


Data Results：

Figure13: Number of colonies



Figure14: CFU / ml