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