Model

Interlab Study

Note

Description: the goal and main contents were quoted from iGEM International InterLab Measurement Study

Methods: the protocol was provided by iGEM InterLab Committee and described briefly in here

Results: the experiment and data presented here were all made by members of team Mingdao

Reference: Fifth International InterLab Measurement Study@iGEM

Instrument

The machine in the Biolab of Mingdao High School: Synergy H1 Hybrid Multi-Mode Microplate Reader

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."

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."

"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?"

Calibration Reference

Calibration 1: OD600 Reference point - LUDOX Protocol

Materials

1ml LUDOX CL-X

ddH2O

96 well Black Clear Bottom Plate

Method

↓ Add 100 μl LUDOX into wells A1, B1, C1, D1

↓ Add 100 μl of ddH2O into wells A2,B2,C2,D2

↓ Measure absorbance at 600 nm

↓ Record the data

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

Materials

300 μL silica beads Microsphere suspension

ddH2O

96 well Black Clear Bottom Plates

Method

↓ Obtain Silica Beads in the InterLab Kit

↓ Pipet 96 μL beads into an eppendorf

↓ Add 904 μL of ddH2O to the microspheres

↓ Vortex well to obtain stock Microsphere Solution.

↓ Preparation of microsphere serial dilutions as follows

↓ Measure Abs 600

↓ Record the data

Result

Raw Data

Particle Standard Curve

Particle Standard Curve(log scale)

Calibration 3: Fluorescence standard curve - Fluorescein Protocol

Materials

Fluorescein

10ml 1xPBS

96 well Black Clear Bottom Plate

Method

Prepare the fluorescein stock solution

↓ Spin down fluorescein tube

↓ Add 1 mL to make 10x fluorescein stock solution (100 μM) of 1xPBS.

↓ Dilute 100 μL of 10x fluorescein stock into 900 μL 1xPBS

↓ Prepare the serial dilutions of fluorescein as follows

↓ Measure fluorescence at Ex/Em = 485/528 nm

Result

Raw Data

Fluorescein Standard Curves

Fluorescein Standard Curves(log scale)

Cell Measurement

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 Black Clear Bottom Plate

Workflow

Method

Day1

↓ transform Escherichia coli DH5 with these plasmids

Day2

↓ Pick 2 colonies from each of the transformed E. coli

↓ inoculate in 5-10 mL LB medium + Chloramphenicol.

↓ Grow the cells overnight (16-18 hours) at 37°C and shake at 220 rpm.

Day 3

↓ Make a 1:10 dilution by 0.5mL of overnight culture into 4.5mL of LB + Chlor

↓ Measure Abs 600 of these 1:10 diluted cultures

↓ Record the data

↓ Dilute the cultures further to Abs600=0.02 in 12ml of media

↓ Shake in a 37°C incubator at 220 rpm for 6 hours.

↓ Measure Abs600 and fluorescence

↓ Record data

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