Team:Manchester/InterLab

Fluorescence assays are amongst the most important measurement tools in biological experiments. However, the direct comparison of fluorescence data is problematic as data is often reported in different units or processed in different ways. This inability to directly compare results makes it harder to work collaboratively between laboratories and thus hinders the advancement of research. Therefore, it is important that a standardised protocol be established, so that groups around the world can achieve better, more streamlined fluorescence measurements. The Fourth International Laboratory Measurement Study aims to test the effectiveness of this standardised protocol by answering the important question: How close can the numbers be when fluorescence is measured all around the world using the same exact protocol?

Manchester iGEM 2017 is proud to have taken part in the Interlab study for the first time. We hope our participation will help benefit other iGEM teams and the synthetic biology community as a whole.
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Protocols

Aseptic technique was maintained at all times.

Transformation

The E.coli strain used was Dh5α. The DNA provided in the distribution kit for the transformations was re-suspended using 10 μl of dH2O. Competent cells were transformed using the following 8 devices, inside the plasmid backbone pSB1C3:

- BBa_R0040 (Negative Control)

- BBa_I20270 (Positive Control)

- BBa_J364000 (Device 1)

- BBa_J364001 (Device 2)

- BBa_J364002 (Device 3)

- BBa_J364003 (Device 4)

- BBa_J364004 (Device 5)

- BBa_J364005 (Device 6)

Transformations were done using our refined protocol (see additional work) and the transformed cells were subsequently plated on 8 LB agar plates containing chloramphenicol at a concentration of 25 mg/ml. The plates were incubated overnight at 37°C.

Overnight Cultures

5 ml of sterile LB broth was transferred to a total of 16 x 50 ml falcon tubes; chloramphenicol was then added to each tube at a concentration of 25 mg/ml. Two colonies were picked from each plate, each individual colony was transferred to a separate falcon tube. The tubes were incubated overnight at 37°C and 180 rpm.

Plate Reader Settings

Measurements of optical density at 600 nm (OD600) and fluorescence were taken using a BMG LabTech Clariostar plate reader, provided to us by the Manchester Institute of Biotechnology (MIB). OD600 measurements were all taken at 24°C with 35 flashes, and an orbital averaging of 3. Fluorescence measurements were taken with 8 flashes and a gain of 748. Excitation and emission wavelengths were 515-20 nm and 470-15 nm respectively. Pathlength correction was turned off for both measurements.

OD600 Reference Point

LUDOX S-40 was used as a single reference point to calibrate the plate reader. A ratiometric conversion factor was obtained to ensure that all Absorbance measurements at 600 nm (Abs600) were converted to OD600 measurements, whilst also taking into account differences in instruments.

100 µl of LUDOX was transferred into wells A1-D1 of a black, clear-bottom 96-well plate. 100 µl of distilled H₂O was then added into wells A2-D2. Abs600 measurements of LUDOX and H₂0 were then taken. A ratiometric conversion factor of roughly 3.269 was obtained.

Fluorescence Standard Curve

The fluorescein stock was spun down for 30 seconds at 3000 rpm and then re-suspended in 1 mL of 1x PBS to produce a 2x stock solution (100 μM). This was then further diluted with 1x PBS to a 1x stock solution with a final concentration of 50 μM.

200 µl of the fluorescein stock was transferred into wells A1-D1. The fluorescein stock was then serially diluted in four replicates (A2-A12, B2-B12, C2-C12, D2-D12) by mixing with 1x PBS to obtain 100 µl of 25, 12.5, 6.25, 3.125, 1.5625, 0.78125, 0.390625, 0.1953125, 0.09765625, 0.048828125, and 0 µM of fluorescein solution. Fluorescence was then measured using a BMG LabTech ClarioStar plate reader, and a fluorescence standard curve generated (Figure 1). This standard curve was used to correct cell based readings to an equivalent fluorescein concentration and measure the concentration of GFP.

Cell Measurements

Overnight cultures of each device were diluted to a target OD of 0.02 and incubated at 37°C and 220 rpm, falcon tubes were wrapped in tin foil throughout. 500 μL of each device was transferred to an Eppendorf tube and put on ice at time points 0 h, 2 h, 4 h, and 6 h. 100 μl of each sample was then transferred to 96 well plate using the provided set-up guidelines and measurements. OD and fluorescence measurements were taken simultaneously.
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Results

Overall, the results of the experiment was unexpected. Out of all the devices tested, only device 2 exhibited the expected trend, the fluorescence of cells increasing with time. Device 4 and the positive control also display this trend up to 4 h before decreasing. However, this decrease could be explained by our cells dying after 6 hours. The negative control displays fluorescence, which was not supposed to happen. We suspect that this may be due to contamination or autofluorescence of colony 2. Colony 1, which was not contaminated, showed very low levels of fluorescence, similar to those seen in device 3 and 6 (See Full Results).

We removed the 4 h time point for device 1 from figure 2. This particular time point exhibited high fluorescence at low OD, and significantly changed the scale of the graph . We suspect that this was potentially due to human error in loading the media.

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Improvements

We are aware that contamination has occurred in colony 2 (Figure 3), as fluorescence was detected in the negative control (which does not have a GFP construct). This is most likely due to human error when transferring samples.

Additional work

We wanted to take full advantage of the Interlab project and use it to improve upon the skills that we have gained since beginning the experimental part of our project. For this reason, we decided to make our own DH5α competent cells. This also gave the rest of the team plenty of stock to work with whilst they continued to complete their own experiments. We used the Benchling protocol which can be found here: https://benchling.com/protocols/PqDdikG7/tss-competent-e-coli-preparation

Transformation

Initially we struggled to obtain colonies from the transformation of test kit 7, refining our protocol led us to successfully transform kit 6. This protocol has since been adopted by other members of our team to improve the efficiency of their own transformations. Our protocol is as follows:

- 2 μl of DNA was added to 100 μl of competent DH5α E.coli cells.

- Cells were incubated on ice for 30 minutes.

- Cells were heat shocked in water bath set at 42°C for 30 seconds.

- 800 μl of SOC media was added and cells were incubated for 60 minutes at 37°C.

- Cells were then spun down in a tabletop centrifuge for 2 minutes at 5000 rpm.

- 600 μl of the supernatant was removed and the pellet re-suspended in the remaining supernatant.

- The remaining 200 μl of re-suspended cells were then plated.

Miniprep and restriction digest

As we initially struggled to transform all the devices in kit plate 7, transformations which yielded colonies were miniprepped and the purity and concentration of the DNA was assessed using a nanodrop spectrophotometer. We then performed a restriction digest (using XbaI and PstI) and ran the DNA on an agarose gel. This confirmed that our transformations had not been successful in all cases, as the results did not match the band size that we predicted using SnapGene. Conversely, the successful transformation of all devices from kit plate 6 was confirmed using the same technique. At this point we then prepared overnight cultures ready for the plate reader measurement Interlab protocol.
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