Team:Mingdao/InterLab
- EXPERIMENT
- Experiment
- MODELING
- Modeling
Interlab Study
Introduction
Reliable and repeatable measurement is a key component to all engineering disciplines. The same holds true for synthetic biology, which has also been called engineering biology. However, the ability to repeat measurements in different labs has been difficult. The Measurement Committee, through the InterLab study, has been developing a robust measurement procedure for green fluorescent protein (GFP) over the last several years. We chose GFP as the measurement marker for this study since it's one of the most used markers in synthetic biology and, as a result, most laboratories are equipped to measure this protein.
The aim to improve the measurement tools available to both the iGEM community and the synthetic biology community as a whole. One of the big challenges in synthetic biology measurement has been that fluorescence data usually cannot be compared because it has been reported in different units or because different groups process data in different ways. Many have tried to work around this using “relative expression” comparisons; however, being unable to directly compare measurements makes it harder to debug engineered biological constructs, harder to effectively share constructs between labs, and harder even to just interpret your experimental controls.
The InterLab protocol aims to address these issues by providing researchers with a detailed
protocol and data analysis form that yields absolute units for measurements of GFP in a plate
reader.
Goal for the Fifth InterLab
The goal of the iGEM InterLab Study is to identify and correct the sources of systematic variability in synthetic biology measurements, so that eventually, measurements that are taken in different labs will be no more variable than measurements taken within the same lab. Until we reach this point, synthetic biology will not be able to achieve its full potential as an engineering discipline, as labs will not be able to reliably build upon others’ work.
In the previous interlab studies, it was shown that by measuring GFP expression in absolute fluorescence units calibrated against a known concentration of fluorescent molecule can greatly reduce the variability in measurements between labs. However, when taking bulk measurements of a population of cells (such as with a plate reader), there is still a large source of variability in these measurements: the number of cells in the sample.
Because the fluorescence value measured by a plate reader is an aggregate measurement of an entire population of cells, we need to divide the total fluorescence by the number of cells in order to determine the mean expression level of GFP per cell. Usually this is done by measuring the absorbance of light at 600nm, from which the “optical density (OD)” of the sample is computed as an approximation of the number of cells. OD measurements are subject to high variability between labs, however, and it is unclear how good of an approximation an OD measurement actually is. If a more direct method is used to determine the cell count in each sample, then potentially another source of variability can be removed from the measurements.
This year, teams participating in the interlab study helped iGEM to answer the following question: Can we reduce lab-to-lab variability in fluorescence measurements by normalizing to absolute cell count or colony-forming units (CFUs) instead of OD?
In order to compute the cell count in the different teams samples, two orthogonal approaches were be used:
1. Converting between absorbance of cells to absorbance of a known concentration of beads.
Absorbance measurements use the way that a sample of cells in liquid scatter light in order to approximate the concentration of cells in the sample. In this year’s Measurement Kit, teams were provided with a sample containing silica beads that are roughly the same size and shape as a typical E. coli cell, so that it should scatter light in a similar way. Because the concentration of the beads is known, each lab’s absorbance measurements can be converted into a universal, standard “equivalent concentration of beads” measurement.
2. Counting colony-forming units (CFUs) from the sample.
A simple way to determine the number of cells in a sample of liquid media is to pour some out on a plate and see how many colonies grow on the plate. Since each colony begins as a single cell (for cells that do not stick together), we can determine how many live cells were in the volume of media that we plated out and obtain a cell concentration for our sample as a whole. Each team will have to determine the number of CFUs in positive and negative control samples in order to compute a conversion factor from absorbance to CFU.
By using these two approaches, Interlab Measurement Study will be able to determine how much they agree with each other, and whether using one (or both) can help to reduce lab-to-lab variability in measurements. If it can, then together we will have brought synthetic biology one step closer to becoming a true, reliable engineering discipline.
Calibration Reference
Calibration 1:OD600 Reference point - LUDOX Protocol
LUDOX CL-X (45% colloidal silica suspension) was used as a single point reference to obtain a conversion factor to transform our absorbance (Abs600) data from our plate reader into a comparable OD600 measurement as would be obtained in a spectrophotometer. Such conversion is necessary because plate reader measurements of absorbance are volume dependent; the depth of the fluid in the well defines the path length of the light passing through the sample, which can vary slightly from well to well. In a standard spectrophotometer, the path length is fixed and is defined by the width of the cuvette, which is constant. Therefore this conversion calculation can transform Abs600 measurements from a plate reader (i.e., absorbance at 600nm, the basic output of most instruments) into comparable OD600 measurements. The LUDOX solution is only weakly scattering and so will give a low absorbance value.
[ IMPORTANT NOTE : many plate readers have an automatic path length correction feature. This adjustment compromises the accuracy of measurement in highly light scattering solutions, such as dense cultures of cells. YOU MUST THEREFORE TURN OFF PATHLENGTH CORRECTION if it can be disabled on your instrument . Our Instrument did not have any pathlength correction].
Materials
1ml LUDOX CL-X (provided in kit)
ddH2 0 (provided by team)
96 well plate, black with clear flat bottom preferred (provided by team)
Method
Add 100 μl LUDOX into wells A1, B1, C1, D1
Add 100 μl of ddH2 O into wells A2,B2,C2,D2
Measure absorbance at 600 nm of all samples in the measurement mode you plan to use for cell measurements
Record the data in the table below or in your notebook
Import data into Excel sheet provided ( OD600 reference point tab )
Result
The table shows the OD600 measured by a spectrophotometer (see table above) and plate reader data for H2O and LUDOX corresponding to the expected results. The corrected Abs600 is calculated by subtracting the mean H2O reading. The reference OD600 is defined as that measured by the reference spectrophotometer. The correction factor to convert measured Abs600 to OD600 is thus the reference OD600 divided by Abs600. All cell density readings using this instrument with the same settings and volume can be converted to OD600 by multiplying by 4.200.
Calibration 2: Particle Standard Curve - Microsphere Protocol
We prepared a dilution series of monodisperse silica microspheres and measured the Abs600 in our plate reader. The size and optical characteristics of these microspheres are similar to cells, and there is a known amount of particles per volume. This measurement allows us to construct a standard curve of particle concentration which can be used to convert Abs600 measurements to an estimated number of cells.
Materials
300 μL silica beadsMicrosphere suspension (provided in kit, 4.7*108 microspheres)
ddH2O (provided by EPFL)
96 well plates, black with clear flat bottom (provided by team)
Method
Preparation of the Microsphere stock solution:
Obtain the tube labeled “Silica Beads” from the InterLab test kit and vortex 4 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).
Immediately pipet 96 μL eppendorf
Add 904 μL of ddH2O to the microspheres
Vortex well to obtain stock Microsphere Solution.
Vortex well to obtain stock Microsphere Solution. Preparation of microsphere serial dilutions:
Accurate pipetting is essential. Serial dilutions will be performed across columns 1-11. COLUMN 12 MUST CONTAIN ddH2O ONLY. Initially you will setup the plate with the microsphere stock solution in column 1 and an equal volume of 1x ddH2O in columns 2 to 12. You will perform a serial dilution by consecutively transferring 100 μL from column to column with good mixing.
1. Add 100 μl of ddH2O into wells A2, B2, C2, D2....A12, B12, C12, D12
2. Vortex the tube containing the stock solution of microspheres vigorously for 10 seconds
3. Immediately add 200 μl of microspheres stock solution into A1
4. Transfer 100 μl of microsphere stock solution from A1 into A2.
5. Mix A2 by pipetting up and down 3x and transfer 100 μl into A3
6. Mix A3 by pipetting up and down 3x and transfer 100 μl into A4...
7. Mix A4 by pipetting up and down 3x and transfer 100 μl into A5...
8. Mix A5 by pipetting up and down 3x and transfer 100 μl into A6...
9. Mix A6 by pipetting up and down 3x and transfer 100 μl into A7...
10. Mix A7 by pipetting up and down 3x and transfer 100 μl into A8...
11. Mix A8 by pipetting up and down 3x and transfer 100 μl into A9...
12. Mix A9 by pipetting up and down 3x and transfer 100 μl into A10...
13. Mix A10 by pipetting up and down 3x and transfer 100 μl into A11...
14. Mix A11 by pipetting up and down 3x and transfer 100 μl into liquid waste TAKE CARE NOT TO CONTINUE SERIAL DILUTION INTO COLUMN 12.
15. IMPORTANT ! Re-Mix (Pipette up and down) each row of your plate immediately before putting in the plate reader! (This is important because the beads begin to settle to the bottom of the wells within about 10 minutes, which will affect the measurements.) Take care to mix gently and avoid creating bubbles on the surface of the liquid.
16. Measure Abs 600 of all samples in instrument
17. Record the data in your notebook
18. Import data into Excel sheet provided ( particle standard curve tab )
Result
Raw Data
Particle Standard Curve
Particle Standard Curve(log scale)
Calibration 3: Fluorescence standard curve - Fluorescein Protocol
Plate readers report fluorescence values in arbitrary units that vary widely from instrument to instrument. Therefore absolute fluorescence values cannot be directly compared from one instrument to another. In order to compare fluorescence output of test devices between teams, it is necessary for each team to create a standard fluorescence curve. Although distribution of a known concentration of GFP protein would be an ideal way to standardize the amount of GFP fluorescence in E. coli cells, the stability of the protein and the high cost of its purification are problematic. The Interlab Study therefore uses the small molecule fluorescein, which has similar excitation and emission properties to GFP, but is cost-effective and easy to prepare. (The version of GFP used in the devices, GFP mut3b, has an excitation maximum at 501 nm and an emission maximum at 511 nm; fluorescein has an excitation maximum at 494 nm and an emission maximum at 525nm).
Teams will prepare a dilution series of fluorescein in four replicates and measure the fluorescence in a 96 well plate in your plate reader. By measuring these in the plate reader, a standard curve of fluorescence for fluorescein concentration will be generated. THus, different teams will be able to use this to convert their cell based readings to an equivalent fluorescein concentration. Before beginning this protocol, teams should ensure that they are familiar with the GFP settings and measurement modes of their instrument. Each team needs to know what filters your instrument has for measuring GFP, including information about the bandpass width (530 nm / 30 nm bandpass, 25-30nm width is recommended), excitation (485 nm is recommended) and emission (520-530 nm is recommended) of this filter.
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)
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 (100 μM) by resuspending fluorescein in 1 mL 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
Prepare the serial dilutions of fluorescein
Accurate pipetting is essential. Serial dilutions will be performed across columns 1-11. COLUMN 12 MUST CONTAIN PBS BUFFER ONLY. Initially you will setup the plate with the fluorescein stock in column 1 and an equal volume of 1xPBS in columns 2 to 12. You will perform a serial dilution by consecutively transferring 100 μl from column to column with good mixing.
1. Add 100 μl of PBS into wells A2, B2, C2, D2....A12, B12, C12, D12
2. Add 200 μl of fluorescein 1x stock solution into A1, B1, C1, D1
3. Transfer 100 μl of fluorescein stock solution from A1 into A2.
4. Mix A2 by pipetting up and down 3x and transfer 100 μl into A3
5. Mix A3 by pipetting up and down 3x and transfer 100 μl into A4...
6.Mix A4 by pipetting up and down 3x and transfer 100 μl into A5...
7.Mix A5 by pipetting up and down 3x and transfer 100 μl into A6...
8.Mix A6 by pipetting up and down 3x and transfer 100 μl into A7...
9. Mix A7 by pipetting up and down 3x and transfer 100 μl into A8...
10. Mix A8 by pipetting up and down 3x and transfer 100 μl into A9...
11. Mix A9 by pipetting up and down 3x and transfer 100 μl into A10...
12. Mix A10 by pipetting up and down 3x and transfer 100 μl into A11...
13. Mix A11 by pipetting up and down 3x and transfer 100 μl into liquid waste TAKE CARE NOT TO CONTINUE SERIAL DILUTION INTO COLUMN 12.
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 )
Result
Raw Data
Fluorescein Standard Curves
Fluorescein Standard Curves(log scale)
Cell Measurement
Prior to performing the cell measurements all three of the calibration measurements should be performed.
For the sake of consistency and reproducibility, Interlab Measurement requires all teams to use E. coli K-12 DH5-alpha.
For all of these cell measurements,we used the same plates and volumes that we used in the calibration protocol.We also used the same settings (e.g., filters or excitation and emission wavelengths) that you used in your calibration measurements.
Materials
Competent cells ( Escherichia coli strain DH5 )
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)
Workflow
Method
Day1
transform Escherichia coli DH5 with these following plasmids (all in pSB1C3):
Thermo-Fisher DH5-alpha Competent Cells (Catalogue #: 18265017 were purchased).
iGEM protocols for resuspending DNA from the kit plates and performing the transformation were used:http://parts.igem.org/Help:Protocols/Transformation
Day2
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.
Day 3
Cell growth, sampling, and assay
Make a 1:10 dilution of each overnight culture in LB+Chloramphenicol (0.5mL of culture into 4.5mL of LB+Chlor)
Measure Abs 600 of these 1:10 diluted cultures
Record the data in your notebook
Dilute the cultures further to a target Abs6 00 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)
Take 500 L 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 500 μl samples per time point, 32 samples total). Place the samples on ice.
Incubate the remainder of the cultures at 37°C and 220 rpm for 6 hours.
Take 500 μl samples of the cultures at 6 hours of incubation into 1.5 ml eppendorf tubes. Place samples on ice.
At the end of sampling point you need to measure your samples (Abs600 and fluorescence measurement), see the below for details.
Record data in your notebook
Import data into Excel sheet provided ( fluorescence measurement tab )
Measurement:
Samples should be laid out according to the plate diagram below. Pipette 100 μl of each sample into each well. From 500 μl 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 100uL each of only LB+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.
Layout for Abs 600 and fluorescence measurement:
Result
Fluorescence Raw Reading
Abs600 Raw Reading
Protocol: Colony Forming Units per 0.1 OD600 E. coli cultures
This procedure was used to calibrate OD600 to colony forming unit (CFU) counts, which are directly relatable to the cell concentration of the culture, i.e. viable cell counts per mL. This protocol assumes that 1 bacterial cell will give rise to 1 colony.
For the CFU protocol, counting colonies is performed for the two Positive Control (BBa_I20270) cultures and the two Negative Control (BBa_R0040) cultures.
Step 1: Starting Sample Preparation
This protocol will result in CFU/mL for 0.1 OD600. Your overnight cultures will have a much higher OD600 and so this section of the protocol, called “Starting Sample Preparation”, will give you the “Starting Sample” with a 0.1 OD600 measurement.
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
LB + Cam before measuring the OD600. Preparation:Add 25 μL culture to 175 μL 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 200 μL .
2.Dilute your overnight culture to OD600 = 0.1 in 1mL of LB + Cam media. Do this in triplicate for each culture.
Use (C1)(V1) = (C2)(V2) to calculate your dilutions
C1 is your starting OD600
C2 is your target OD600 of 0.1
V1 is the unknown volume in μL
V2 is the final volume of 1000 μL
Important:
When calculating C1, subtract the blank from your reading and multiple by the dilution factor you used.
Example: C1 = (1:8 OD600 - blank OD600) x 8 = (0.195 - 0.042) x 8 = 0.153 x 8 = 1.224
Example:
(C1)(V1) = (C2)(V2)
(1.224)(x) = (0.1)(1000μL)
x = 100/1.224 = 82 μL culture
Add 82 μL of culture to 918 μL media for a total volume of 1000 μL
3.Check the OD600 and make sure it is 0.1 (minus the blank measurement). Recommended plate setup is below. Each well should have 200 μL .
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 1900 μL of LB + Cam media for Dilutions 1, 2, and 3 (see figure below).
3. Prepare two 1.5 mL tubes (24 total) with 900 μL 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 100 μL of Starting Culture into Dilution 1.Discard tip.Do NOT pipette up and down. Vortex tube for 5-10 secs.
6. Repeat Step5 for each dilution through to Dilution 5 as shown below.
7. Aseptically spead plate 100 μLon 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
Based on the assumption that 1 bacterial cell gives rise to 1 colony, colony forming units (CFU) per 1mL of an OD600 = 0.1 culture can be calculated as follows:
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.
Example using Dilution 4 from above
# colonies x Final Dilution Factor = CFU/mL
125 x (8 x 105) = 1 x 100000000 CFU ⁄ mL in Starting Sample (OD600 = 0.1)
Result
Colony Forming Units per o.1 OD600 E.coli cultures
