Team:York/Experiments

YORK


Protocols

QWACC Team Methods


In the interest of reproducibility, we would like to use this page to share the protocols that we have used during this project.
This library of protocols has been developed alongside our supervisors, to whom we are very grateful.
Please find, below, the exact methods we've used to generate our data and results.


Growth Testing

E. coli LW06 Medium Growth Test - Maltose

Tris phosphate and maltose concentrations: To determine how LW06 would grow on carbon source maltose (proposed export of Chlamydomonas reinhardtii). Performed in 96 well plates over 24 hours in 96 plate reader. OD reading taken every 30 minutes.

  1. Grow stock culture of LW06 16-24 hours.
  2. Dilute stock culture to produce inoculum with consistent cell density:
    1. Produce inoculum of between 0.5-1.0 OD.
    2. Spin down culture in centrifuge (2500 rpm 2 minutes).
    3. Remove supernatant and suspend in TP medium, repeat twice. (Do not do this for LB control culture.)
  3. Produce TP maltose stock solution:
    1. Make 10 g/L (29.21414 mM) maltose monohydrate solution.
      1. 50 mL distilled water.
      2. 0.5 g maltose.
  4. Make media dilutions in 96 well plates:
    1. Columns 1 & 3 add 200 μL of LB. Columns 2 & 4 add 200 μL of TP. Columns 1 & 2 and columns 3 & 4 will be blanks and growth controls, respectively.
    2. Column 5 add 20 μL of TP.Maltose stock and 180 μL of TP.
    3. Repeat for column 6-12 while increasing TP.Maltose volume by 20 μL for each column. Dilution method results in a 1-8 g/L gradient in the wells 5-12, respectively.
  5. Inoculate plate:
    1. 10 μL of LW06 to columns 3 to 12.
  6. Measure optical density of cultures:
    1. 96 well plate reader, run time 24 hours. One reading per 30 minutes. 48 total readings.

Co-culture ratio experiment

Co-culture experiment: The growth disparity between C. reinhardtii and the E. coli LW06 strain is quite large. LW06 doubles every 28 minutes at 37°C while C. reinhardtii doubles once every 8 hours at 25°C, around 17 times slower than LW06. Considering this, we endeavoured to understand which ratios of initial inoculation would be required to ensure that E. coli does not completely saturate the culture.

  1. Grow stock culture of LW06 16-24 hours.
  2. Dilute stock culture to produce inoculum with consistent cell density:
    1. Produce inoculum of between 0.5-1.0 OD.
    2. Spin down culture in centrifuge (2500 rpm 2 minutes).
    3. Remove supernatant and suspend in TP media, repeat washing step twice.
  3. Grow stock culture of C. reinhardtii 7 days minimum (need long term stock culture from which to draw):
    1. Produce inoculum of 10 mL of between 0.5-1.0 OD.
    2. Spin culture down in centrifuge (2500 rpm 2 minutes.
    3. Remove supernatant and suspend in TP, repeat washing step twice.
  4. Produce TP maltose and TAP maltose stock media:
    1. Make 10 g/L (29.21414 mM) of TP.Maltose:
      1. 50 mL TP in each tube.
      2. 0.50 g maltose in each tube.
    2. Make 10 g/L (29.21414 mM) of TAP.Maltose:
      1. 50 mL TAP in each tube.
      2. 0.50 g maltose in each tube.
  5. Fill 8 falcon tubes (50 mL) in the following manner:
    1. Tube 1 TP 25 mL.
    2. Tube 2 TP 25 mL.
    3. Tube 3 TAP 25 mL.
    4. Tube 4 TAP 25 mL.
    5. Tube 5 TP.M 25 mL.
    6. Tube 6 TP.M 25 mL.
    7. Tube 7 TAP.M 25 mL.
    8. Tube 8 TAP.M 25 mL.
  6. Inoculate all tubes with 100 μL of C. reinhardtii. Inoculate the first tube in each pair with 10 μL and the second tube with 100 μL of LW06.
  7. Leave to grow for 8 days in growth room at 25°C (we used an 8 hour day light cycle in the room).
  8. Measure cell types at the end of 8 days via the following methods:
    1. C. reinhardtii haemocytometer microscope counting at a 104 dilution.
    2. E. coli LW06 LB spread plating at a 104 dilution.

LW06 Medium Growth Test - Hblend

Hblend (Home blend media) experiment: TP media is quite depleted in most micronutrients required for E. coli growth, as it is a media formulated for C.reinhardtii growth under photosynthetic growth. So, to try and solve this problem we decided to create a growth medium that could allow for the improved growth of LW06 and C. reinhardtii. Hblend media like M9 medium will act as a micronutrient supply with a variable carbon source. This test was performed in 96 well plates over 24 hours in 96 plate reader with an OD reading taken every 30 minutes.

  1. Grow stock culture of LW06 16-24 hours.
  2. Dilute stock culture to produce inoculum with consistent cell density:
    1. Produce inoculum of between 0.5-1.0 OD.
    2. Spin down culture in centrifuge (2500 rpm 2 minutes).
    3. Remove supernatant and suspend in TP and Hblend media, repeat twice. (Do not do this for LB control cuture.)
  3. Produce Home blend medium stock (500 mL):
    1. Prepare micronutrients:
      1. 1 M NaCl: 4.28 mL
      2. 1 M Na2HPO4: 23.875 mL
      3. 1 M KH2PO4: 11.02 mL
      4. 1 M NH4Cl: 9.345 mL
      5. 0.1 M CaCl2: 1.7 mL
      6. 1 M MgSO4: 1 mL
      7. 28.5 μM (NH4)6Mo7O24: 0.5 mL
      8. 0.1 mM Na2SeO3: 0.5 mL
      9. 2.5 mM Zn x EDTA: 0.5 mL
      10. 6 mM Mn x EDTA: 0.5 mL
      11. 20 mM Fe x EDTA: 0.5 mL
      12. 2 mM Cu x EDTA: 0.5 mL
      13. Phosphate solution: 0.1875 mL
      14. 2M Tris/acetate or tris/HCl, pH 7.4: 5 mL
    2. Add distilled water 440.6 mL to make 500 mL.
    3. Autoclave to sterilize.
    4. Add 20 mL 20% glucose solution or other carbon source.
  4. Set up 96 well plates:
    1. Columns 1-6 add 200 μL of LB, TP.Maltose, TP.Glucose, Hblend, Hblend.Glucose and Hblend.Maltose, respectively.
    2. Columns 7-12 add 190 μL of LB, TP.Maltose, TP.Glucose, Hblend, Hblend.Glucose and Hblend.Maltose, respectively.
  5. Inoculate plate:
    1. 10 μL of LW06 to each of columns 7-12 respective to washed down inoculating culture.
  6. Measure optical density of cultures:
    1. 96 well plate reader, run time 24 hours. One reading per 30 minutes, 48 readings total.

C. reinhardtii Ethanol Tolerance

Experiment: The key to a self-sustaining (or even partially self-sustained) biological production culture is that the carbon provider (C. reinhardtii) is not killed by the product (ethanol) produced by the other organism (LW06) in the culture. So, we set up an ethanol tolerance test for C. reinhardtii in a 24 well plate.

  1. Grow stock culture of C. reinhardtii 7 days minimum (need long term stock culture from which to draw):
    1. Produce inoculum of 10ml of between 0.5-1.0 OD.
    2. Spin culture down in centrifuge (2000 rpm 2 minutes).
    3. Remove supernatant and suspend in TP, repeat washing step twice.
  2. Make media dilutions in 24 well plates: one plate for TP medium, one for TAP medium.
    1. Plate 1 TAP:
      1. Rows A-C TAP.
      2. Row D distilled water.
      3. Column 1 leave un-inoculated for all rows. A-C 1000 μL TAP; D 1000 μL distilled water.
      4. Column 2 rows A-C 1000 μL TAP; row D 1000 μL distilled water. Inoculate with 250 μL of C. reinhardtii.
      5. Column 3 rows A-C 990 μL TAP, 10 μL 100% ethanol; row D 990 μL distilled water, 10 μL 100% ethanol.
      6. Repeat above for columns 4-6, increasing volume of ethanol and decreasing TAP/distilled water volumes in increments of 10 μL.
      7. Ethanol gradient increases from 1% to 4% from column 3 to column 6, respectively.
    2. Plate 2 TP:
      1. Repeat above method, replacing TAP with TP.
  3. Inoculate plate:
    1. 250 μL of C. reinhardtii inoculum in columns 2-6.
  4. Measure optical density of cultures:
    1. 24 well plate reader (λ = 720 nm). Readings taken once ~24 hours. When not being measured, keep in 25°C incubator room/cabinet with 8 hour day cycle.

DIHM & Milli-fluidic Device Methodology

Milli-fluidic Chamber Casting from PDMS

Crating chamber for imaging microorganisms in liquid phase using transparent non-toxic polymer PDMS.

  1. Gather materials:
    1. PDMS
    2. PDMS curing agent
    3. Falcon tube 20 ml
    4. Iso-propyl alcohol
    5. Magic tape
    6. Spatula
    7. Milli-fluidic Chamber mould
    8. Pipettes
  2. Clean mould
    1. Rinse mould with Iso-propyl alcohol and then with deionised water. Left drying.
    2. If there some pieces of rubbish on the mould use magic tape to remove them.
  3. Curing PDMS
    1. Pour 4 ml of PDMS and 0.4 ml of curing agent into falcon tube and mix well with disposable stick or metallic spatula.
    2. Put falcon tube with PDMS into centrifuge. Use another falcon tube with water as counterweight. Spin for 2 min at 2500 rpm.
  4. Moulding
    1. Take falcon tube with PDMS out of centrifuge. It should not contain any bubbles. Pour it into the mould.
    2. Put mould with PDMS into the oven at 80 Centigrade for 2 hours. If it is not enough, leave in oven for longer until PDMS fully solidifies.
    3. Take mould with PDMS out of the oven and let it cool for a minute. Take chamber out with spatula.

Milli-fluidic Chamber Production

Low cost chamber for imaging microorganisms

  1. Gather materials:
    1. Acrylic sheet
    2. Transparent tubing d = 1.5 mm
    3. Epoxy glue
    4. Glass slide and coverslip
    5. Blu tack
  2. Design
    1. Chamber design will depend on what type of microorganisms you are working and imaging technique you are using. Tubing has to be taken into account as well.
    2. Main parameters for our chamber:
      • Height should be 2 mm - 5 mm. Physical limitation of laser we are using.
      • We want channe to be l as narrow as possible. It will match diameter of tubing.
      • No sharp edges. This will prevent algae from sticking in chamber.
      • Cheap and easy to produce.
  3. Drafting
    1. Draw your chamber on paper and then transfer it into 3D CAD specifying all dimensions. We were using Fusion 360. It is free for students.
    2. Engraving for tubing is a tricky part. It is unique for every laser/material combination. You will have to play around with settings to achieve deepness that you want.
    3. All laser cutting was done by a trained specialist.
    4. It is important to have 3D model of the chamber. All machines are different and do not support same type of files, but having one file that illustrates your design in detail will help to transfer it onto laser cutter.
  4. Manufacturing
    1. Laser cut chamber from acrylic sheet.
    2. It will take multiple attempts to make right engraving.
  5. Tubing
    1. Cut enough tubing for peristaltic pump. Two tubes 30 cm long should be enough.
    2. Apply super or epoxy glue around one end of each tube and put it into engraved part of the chamber. If necessary, add some glue from sides were tubing sticks out.
    3. Put glued chamber into oven to accelerate process. 4 hours at 60 Centigrade is enough for epoxy glue.
  6. Assembly
    1. Most innovative part of the project.In order to stick glass slide and coverslip to chamber we are using blu tack.
    2. Create thin rolls around 0.5 mm in diameter out of blu tack.
    3. Make an outline around channel from both sides.
    4. Squeeze chamber between slide and cover slip.

Diffraction Pattern Imaging with Raspberry Pi

Imaging diffraction patterns cheaply: in order to keep the DIHM cheap, we planned to use a Raspberry Pi 3 and Camera Module V2 as the image capturing device. We tested its capabilities on an optical bench.

  1. Gather materials:
    1. Raspberry Pi 3 and power supply.
    2. Camera Module V2 (remove lens to expose CMOS).
    3. HDMI cable.
    4. USB keyboard.
    5. USB mouse.
    6. Monitor (with HDMI input).
    7. Optical bench (and clamps, stands, related accessories).
    8. Laser (we used a 635 nm laser diode with collimating lens at this stage, though the collimating lens was omitted in later tests due to its projection of concentric rings into images).
    9. Glass slides and cover slips.
    10. Microbeads of known diameter.
  2. Attach Camera Module V2 to Raspberry Pi.
  3. Set up Raspberry Pi 3 for ease of control:
    1. Connect HDMI from Pi to monitor.
    2. Connect mouse and keyboard to the Pi.
    3. Connect power supply to Pi and switch on Pi and monitor.
  4. Place the Camera Module and laser at opposite ends of the optical bench.
  5. Align the laser with the Camera Module's CMOS:
    1. Place a piece of card in front of the laser, in a clamp, on the optical bench.
    2. Make a mark where the laser meets the card.
    3. Move the card along the optical bench, towards the camera.
    4. Adjust the laser and repeat until it follows the mark on the card all the way along the bench.
    5. Ensure that the Camera Module is placed directly in the path of the laser now that it is aligned with the bench.
  6. Add a suitably diluted (trial and error) sample of microbeads to a slide.
  7. Place the slide (in a clamp) on the optical bench, as close to the CMOS as possible.
  8. Take images of the microbeads' diffraction pattern:
    1. Open the terminal (command prompt) on the Pi's interface.
    2. Use the raspistill command to capture a monochrome image:
      • raspistill -e png -cfx 128:128 -w 512 -h 512 -o /home/pi/ExampleFilename.png
      • The above exemplar code takes a monochrome, 512 x 512 pixel image via the Camera Module and saves the picture in PNG format in the /home/pi/ folder under the filename "ExampleFilename.png".
      • Taking pictures in monochrome prevents the Pi from trying to colour balance the images, which can alter the clarity, while keeping it to 512 x 512 pixels allows for quick data processing in later stages. PNG format is used as it is lossless, preserving all quality.
  9. Circular diffraction patterns, caused by the microbeads, should be apparent in the captured picture.

Organism Image Capturing on Glass Slides, Mouse and Keyboard

Taking images for hologram production, using glass slides and a mouse and keyboard. Our original plan, to use a flow chamber to cycle through liquid cultures/co-cultures in a closed system, needed improvement. Hence, we resorted to using glass slides. Further, our control software had bugs that prevented us from using it, so we used a mouse, keyboard and python programs to manually capture images.

At least 20 images are required to make an appropriate background image from which holograms can be created, so we tried to take between 20 and 50 images of each sample. We have written software to make this easier (see Downloads: "Taking Series of Images (Python)"). This protocol assumes that it is stored on the Pi from the beginning.

  1. Gather materials:
    1. Constructed DIHM (as per the Hardware page).
    2. Organism of choice.
    3. TAP medium.
    4. HDMI cable.
    5. USB keyboard.
    6. USB mouse.
    7. Monitor (with HDMI input).
    8. Glass slides and cover slips.
    9. Size 4 Allen key.
    10. Small spirit level(s).
  2. Arrange platforms for desired magnification:
    1. Use Allen key to loosen/tighten into place.
    2. Use spirit level to ensure lens platform is level with base of DIHM.
      • For our best results, we used a sample to lens distance of around 1.2 cm (placing slides directly on top of lens platform) and a lens to camera distance of 6 cm, giving a magnification of 5x.
  3. Set up Pi connections:
    1. Connect HDMI, USB mouse and USB keyboard to Pi through base of DIHM.
    2. Connect HDMI to monitor, also.
    3. Plug in Pi's power supply to mains voltage and switch on. (Laser will become active.)
  4. Prepare 10-12 µL of suitably TAP-diluted (trial and error) sample of the organism on a glass slide.
  5. Place the slide at the desired height on the DIHM.
  6. Take images:
    1. Begin moving the slide across the camera's field of view, slowly.
    2. Open the terminal (command prompt) on the Pi's interface.
    3. While moving the slide, use the python program to capture 50 images:
      • python /home/pi/Take50Images.py
      • Try to avoid passing the edges of the cover slip while taking images, as these pictures will be averaged to create a background. This can then be used to create holograms.
  7. 50 images should now be saved in the folder "/home/pi/DATE--ORGANISM--DILUTION--MAGNIFICATION/".

Organism Image Capturing with a Milli-fluidic Chamber, Mouse and Keyboard

Taking images for hologram production, using a milli-fluidic chamber and a mouse and keyboard. We connected the chamber to liquid culture in a closed system. Our control software had bugs that prevented us from using it, so we used a mouse, keyboard and python programs to manually capture images.

At least 20 images are required to make an appropriate background image from which holograms can be created, so we tried to take between 20 and 50 images of each sample. We have written software to make this easier (see Downloads: "Taking Series of Images (Python)"). This protocol assumes that it is stored on the Pi from the beginning.

  1. Gather materials:
    1. Constructed DIHM (as per the Hardware page).
    2. Constructed milli-fluidic chamber (as per protocol).
    3. Peristaltic pump with flow-rate on order millilitres.
    4. Organism of choice in liquid culture.
    5. TAP medium.
    6. 100% ethanol.
    7. HDMI cable.
    8. USB keyboard.
    9. USB mouse.
    10. Monitor (with HDMI input).
    11. Size 4 Allen key.
    12. Small spirit level(s).
  2. Arrange platforms for desired magnification:
    1. Use Allen key to loosen/tighten into place.
    2. Use spirit level to ensure lens platform is level with base of DIHM.
      • For our best results, we used a sample to lens distance of around 1.2 cm (placing slides directly on top of lens platform) and a lens to camera distance of 6 cm, giving a magnification of 5x.
      • However, when we used the milli-fluidic chambers, we hadn't yet built the lens platform. So, we placed the chambers directly over the Pi's Camera Module, giving an effective magnification of 1x.
  3. Set up Pi connections:
    1. Connect HDMI, USB mouse and USB keyboard to Pi through base of DIHM.
    2. Connect HDMI to monitor, also.
    3. Plug in Pi's power supply to mains voltage and switch on. (Laser will become active.)
  4. Clean the milli-fluidic chamber:
    1. Pump 100% ethanol through the chamber to sterilise it.
    2. Pump TAP medium through the chamber to ensure that no ethanol remains.
  5. Connect the chamber and liquid culture to the pump.
  6. Place the chamber at the desired height on the DIHM.
  7. Take images:
    1. Switch on the pump.
    2. Open the terminal (command prompt) on the Pi's interface.
    3. Ensure the chamber is in the camera's field of view, then use the python program to capture 50 images:
      • python /home/pi/Take50Images.py
  8. 50 images should now be saved in the folder "/home/pi/DATE--ORGANISM--DILUTION--MAGNIFICATION/".

Organism Image Capturing with a Milli-fluidic Chamber and Control Sotware

Taking images for hologram production, using a milli-fluidic chamber and control software. We connected the chamber to liquid culture in a closed system. Our control software had bugs that prevented us from using it during the testing phase, so we have written a protocol detailing how we would have used it, had we been able to.

  1. Gather materials:
    1. Constructed DIHM (as per the Hardware page).
    2. Constructed milli-fluidic chamber (as per protocol).
    3. Peristaltic pump with flow-rate on order millilitres.
    4. Organism of choice in suitably diluted liquid culture.
    5. TAP medium.
    6. 100% ethanol.
    7. Computer capable of running control software.
    8. Ethernet cable.
    9. Size 4 Allen key.
    10. Small spirit level(s).
  2. Arrange platforms for desired magnification:
    1. Use Allen key to loosen/tighten into place.
    2. Use spirit level to ensure lens platform is level with base of DIHM.
  3. Set up Pi connections:
    1. Connect ethernet cable from control computer to Pi through base of DIHM.
    2. Plug in Pi's power supply to mains voltage and switch on. (Laser will become active.)
  4. Clean the milli-fluidic chamber:
    1. Pump 100% ethanol through the chamber to sterilise it.
    2. Pump TAP medium through the chamber to ensure that no ethanol remains.
  5. Connect the chamber and liquid culture to the pump.
  6. Place the chamber at the desired height on the DIHM.
  7. Take images using the control software:
    1. Turn on the pump.
    2. Ensure the chamber is in the camera's field of view.
    3. Run the control program.
    4. Follow the instructions given within the software.

We intended for the control software to operate as follows:

The control software takes a series of images at regular time intervals, while the culture is pumped through the chamber. It then renders one of the images into a hologram and detects the number of cells of a given size.

DNA Imaging, Analysis and Quantification

1% Agarose Gel for Electrophoresis

  1. Gather materials:
    1. Agarose powder: 0.6 g
    2. TAE (x1): 60 mL
    3. SYBR Safe: 6 μL
    4. Microwave safe flask
    5. Tray for gel casting
    6. Well comb
  2. Mix Agarose powder and TAE into microwaveable flask.
  3. Microwave for 1-3 min until agarose fully dissolved:
    1. Microwave in 30s pulses, swirling the flask occasionally as the solution heats up.
    2. Do not overboil the solution, as some of the buffer will evaporate.
  4. Let agarose solution cool down to about 50°C. (Around when you can comfortably keep your hand on the flask.) This should take about 5 minutes.
  5. Add SYBR Safe.
  6. Pour the agarose into a gel tray and place well comb.
    • Different well combs should be used for qualitative or extraction gels.
  7. Place newly poured gel at 4°C for 10-15 minutes OR let sit at room temperature for 20-30 mins, until completely solidified.

Loading and Running Agarose Gel

  1. Gather materials:
    1. 1% agarose gel
    2. TAE buffer (x1)
    3. Loading dye
    4. Ladder
    5. DNA samples/li>
    6. PCR tubes
    7. Electrophoresis unit
    8. Voltage source
  2. Add DNA and loading dye in PCR tube and mix.
    • We use NEB Purple (x6) gel loading dye.
  3. Place gel in electrophoresis chamber.
  4. Fill tank with TAE to submerge gel.
  5. Load 6 μL of ladder and 6 μL of reactions.
    • We use NEB Quick-Load Purple 2-Log DNA Ladder.
    • Other ladders (specified in data analysis) were used when different band size distribution was required.
    • For gel extraction larger sample amounts were loaded.
  6. Run gel at 80 V until bands separated.
  7. Image using UV box.
    • Blue light transilluminator was used to image gels for extraction.

Nanodrop

  1. Add 1.2 µl of pure water to the pedestal and lower the arm.
  2. Initialise the instrument and take blank measurement (as a reference).
  3. Load 1.2 µl of the plasmid prep onto the pedestal.
  4. Load 6 μL of ladder and 6 μL of reactions.
  5. Close the sampling arm and press “Measure” on the computer screen.
  6. Make a note of:
    • The concentration given (ng/µl).
    • The ratio of 260/280 nm. (This should be around 1.9 if the DNA is pure).

Media, Buffers and Solutions

Homeblend Medium Recipe

This recipe is an attempt to create a supplemented TAP/TP to suit E. coli growth, as TAP is highly deficient on several elements.

The recipes for (NH4)6Mo7O24, Na2SeO3, Zn×EDTA, Mn×EDTA, Fe×EDTA, Cu×EDTA, Phosphate solution, tris-acetate, and tris-HCl can be found in the recipe for TAP/TP media.

  1. Medium Recipe, 1 L
    Ingredient Stock Concentration Volume
    NaCl 1 M 8.56 mL
    Na2HPO4 1 M 47.75 mL
    KH2PO4 1 M 22.04 mL
    NH4Cl 1 M 18.69 mL
    CaCl2 0.1 M 3.4 mL
    MgSO4 1 M 2 mL
    (NH4)6Mo7O24 28.5 μM 1 mL
    Na2SeO3 0.1 mM 1 mL
    Zn x EDTA 2.5 mM 1 mL
    Mn x EDTA 6 mM 1 mL
    Fe x EDTA 20 mM 1 mL
    Cu x EDTA 2 mM 1 mL
    Phosphate solution 0.375 mL
    Tris/acetate or tris/HCl, pH 7.4 2 M 10 mL
  2. Add distilled water to bring the volume up to 980 mL.
  3. Autoclave to sterilise.
  4. Add 20 mL 20% glucose solution, or other carbon source of your choice, to bring the solution up to 1 L.

M9 Stock Recipes

    1. M9 Salts, 1 L
      Ingredient Mass
      Na2HPO4.7H2O 64 g
      KH2PO4 15 g
      NaCl 2.5 g
      NH4Cl 5.0 g
    2. Add distilled water to bring the volume up to 1 L.
    3. Autoclave to sterilise.
    1. M9 Medium, 1 L
      Ingredient Volume
      M9 Salts 200 mL
      MgSO4, 1 M 2 ml
      CaCl2, 1 M 100 μl
    2. Add distilled water to bring the volume up to 980 mL.
    3. Autoclave to sterilise.
    4. Add 20 mL 20% glucose, or other carbon source of your choice.

Stock Recipes

    1. TAP Salts, 100 mL
      Ingredient Mass
      CaCl2.2H2O 0.2 g
      MgSO4.7H2O 0.4 g For -S, use 3.3 g MgCl2.6H2O
      NH4Cl 1.5 g
    2. Add distilled water to bring the volume up to 100 mL.
    3. Autoclave to sterilise.
    1. Tris-acetate Solution, 2 M, 100 mL
      Ingredient Amount
      Trizma base 24.2 g
      Glacial acetic acid 10 ml
    2. Add distilled water to bring the volume up to 100 mL.
    3. Filter using 0.2 µm pore OR autoclave using liquid 60 cycle (at least 30 minutes to completely dissolve Tris base) to sterilise.
      • Note: to dissolve Trizma base in a timely manner, stir vigorously and heat gently. It will take some time.
    1. Tris-HCl Solution, 2 M, 100 mL
      Ingredient Amount
      Trizma base 24.2 g
    2. Add around 50 mL distilled water, heat and stir to dissolve.
      • Note: DO NOT add too much water; a substantial amount of HCl will be added in the next step.
      • For the next step, ensure the use of a Tris appropriate pH probe, since some types of probe are not suitable.
    3. Adjust the pH of your solution using 5 M or higher HCl solution.
    4. Add distilled water to bring volume to 100 mL.
    5. Sterilize by filtering using 0.2 µm pore OR autoclave.
    1. Phosphate Solution, 100 mL
      Ingredient Mass
      K2HPO4 28.8 g
      KH2PO4 14.4 g
    2. Add distilled water to bring volume to 100 mL.
    3. Sterilize by filtering using 0.2 µm pore.

Trace Elements

    1. EDTA-Na2
      1. Make concentrated stock solution, 125 mM, 300 mL
        Ingredient Mass
        EDTA-Na2 13.959 g
        • Add to 250 mL distilled water, titrate to pH 8.0 with trace element grade KOH, ~ 1.7 g.
        • Add distilled water to 300 mL.
      2. Make diluted stock solution, 25 mM, 250 mL
        • Add 50 mL of 125 mM concentrated stock to 200 mL distilled water.
    1. (NH4)6Mo7O24
      1. Make concentrated stock solution, 285 μM, 250 mL
        Ingredient Mass
        (NH4)6Mo7O24 0.888 g
        • Add distilled water to 250 mL.
      2. Make diluted stock solution, 28.5 μM, 250 mL
        • Add 25 mL of 280 μM concentrated stock to 225 mL distilled water.
    1. Na2SeO3
      1. Make concentrated stock solution, 1 mM, 250 mL
        Ingredient Mass
        Na2SeO3 0.043 g
        • Add distilled water to 250 mL.
      2. Make diluted stock solution, 0.1 mM, 250 mL
        • Add 25 mL of 1 mM concentrated stock to 225 mL distilled water.
    1. Zn x EDTA
      Ingredient Amount
      ZnSO4.7H2O 0.018 g
      EDTA-Na2, 125 mM 12 mL
      • Add distilled water to 250 mL.
    1. Mn x EDTA
      Ingredient Amount
      MnCl2.4H2O 0.297 g
      EDTA-Na2, 125 mM 12 mL
      • Add distilled water to 250 mL.
    1. Fe x EDTA
      Ingredient Amount
      FeCl3.6H2O 1.35 g
      EDTA-Na2 2.05 g
      Na2CO3 0.58 g
      • Combine EDTA-Na2 and Na2CO3 in water, mix until dissolved.
      • Add FeCl3.6H2O.
      • Add distilled water to bring the volume up to 250 mL.
        • Note: DO NOT use the EDTA-Na2 concentrate for this step.
    1. Cu x EDTA
      Ingredient Amount
      CuCl2.2H2O 0.085 g
      EDTA-Na2, 125 mM 4 mL
      • Add distilled water to 250 mL.

TAP Recipe

    1. TAP, 1 L
      Ingredient Amount
      Tris-acetate, 2 M 10 mL
      TAP salts 25 mL
      Phosphate solution 0.375 mL
      EDTA-Na2, 25 mM 1 mL
      (NH4)6Mo7O24, 28.5 µM 1 mL
      Na2SeO3, 0.1 μM 1 mL
      Zn x EDTA, 2.5 mM 1 mL
      Mn x EDTA, 6 mM 1 mL
      Fe x EDTA, 20 mM 1 mL
      Cu x EDTA, 2mM 1 mL
      Agar (for plates) 15 g
    2. Add to 900 mL distilled water.
    3. Adjust to pH 7.4.
    4. Add distilled water to bring the volume up to 1 L.
    5. Autoclave to sterilise.

TP Recipe

    1. TP, 1 L
      Ingredient Amount
      Tris-HCl, 2 M 10 mL
      TAP salts 25 mL
      Phosphate solution 0.375 mL
      EDTA-Na2, 25 mM 1 mL
      (NH4)6Mo7O24, 28.5 µM 1 mL
      Na2SeO3, 0.1 μM 1 mL
      Zn x EDTA, 2.5 mM 1 mL
      Mn x EDTA, 6 mM 1 mL
      Fe x EDTA, 20 mM 1 mL
      Cu x EDTA, 2mM 1 mL
      Agar (for plates) 15 g
    2. Add to 900 mL distilled water.
    3. Adjust to pH 7.4.
    4. Add distilled water to bring the volume up to 1 L.
    5. Autoclave to sterilise.

TAE Buffer

  1. For 1 L of x50 stock:
    Ingredient Amount
    Tris-base 242 g
    Acetate (100% acetic acid) 57.1 mL
    EDTA (0.5 M sodium EDTA) 100 mL
  2. Add H2O up to 1 L.

LB Medium

  1. For 1 L:
    Ingredient Amount
    Tryptone 10 g
    Yeast Extract 5 g
    NaCl 10 g
    For plates, Agar 12 g
  2. Add to 900 mL H2O
  3. Adjust pH to 7.5 with NaOH
  4. Adjust volume to 1 L with distilled water.

Transformations and Preparation

Chlamydomonas Transformation

  1. Preparing Chlamydomonas batch culture for transformation
    • Inoculation to produce a 4 L culture:
    1. Inoculate 50 mL of TAP with a single, fresh colony from a TAP streak plate.
      • Use a 250 mL conical flask for efficient aeration and mixing while shaking.
    2. Grow, while continuously shaking, for 5 days at 50 – 100 μE until culture density reaches ~ 1 x 107 cells/mL.
    3. Transfer 50 mL into 400 mL of fresh TAP.
    4. Grow while shaking for another 2 days at 50 – 100 μE until culture density reaches ~ 8-9 x 106 cells/mL.
    5. Inoculate 4L of fresh TAP with all 450 mL of Chlamydomonas, grow while bubbling air through and continuously stirring for ~30 hours at 100 μE until culture density reaches 2-4 x 106 cells/mL.
      • Ensure the final inoculation uses an exponential-phase culture.
      • For smaller (up to 1 L) volumes, this step can be done in a conical flask with continuous shaking.
      • The high volume is in case the culture does not reach the desired concentration, in which case a large volume of cells can be spun down for transformation.
  2. Transformation
    • Work with open cultures should be performed in an appropriate flow hood to decrease contamination risk.
    • Use 50 mL Chlamydomonas, at 2-4 x 106 cells/mL, per transformation.
    • Each transformation requires 5 x 107 cells.
    • Materials to prepare
      1. TAP sucrose, 10 mM sucrose – 15 mL per transformation
      2. Selection plates
        • Plate preferences depend on the type of screening to be performed.
        • We used rectangular plates (250 mm x 250 mm).
        • 1.5% TAP agar with appropriate antibiotic.
        • Label on the side to avoid interference with the colony picker camera during selection.
      3. DNA insert cassettes digested with appropriate enzyme
      4. Electroporation cuvettes
      5. Centrifuge tubes
      6. Glass L-shaped spreaders
      7. Pipettes and tips
    • Harvesting
      1. Aliquot Chlamydomonas culture into centrifuge tubes.
      2. Pellet Chlamydomonas culture using centrifuge (4 minutes at 1000x g).
      3. Discard supernatant and avoid disturbing cell pellet.
    • Plating and growth
      1. Use TAP 40 mM sucrose to dilute cell pellet to 2 x 108 cells/mL.
      2. Add 250 μL of Chlamydomonas culture per electroporation cuvette.
      3. Incubate at 16°C fo 5 minutes.
      4. Prepare recovery tubes (15 mL falcon tubes) with 8 mL TAP sucrose.
        • Steps v-vii should be performed as quickly as possible. Especially step vii.
        • Before electroporation, dry the cuvette.
      5. Add transformation cassette DNA (14.5 ng/kb cassette per 250 µL) and mix briefly.
        • Do not mix by repeatedly pipetting. Use the pipette tip to mix by stirring.
      6. Electroporator should be set to 800 V, 25 μF.
      7. Administer pulse and immediately add the mixture from the cuvette to the recovery tube.
    • Electroporation
      1. After all electroporations are complete, transfer recovery tubes to a shaker, and grow for 6 hours in the dark.
      2. Centrifuge the recovery tubes for 4 min at 1000x g.
      3. Discard most of the supernatant, leaving about 500 µL.
      4. Resuspend the pellet in the residual supernatant and spread on a transformation plate using a glass spreader.
      5. After all plates are adequately dry, place them upside-down in stacks in plastic bags, and return to growth room.
      6. Incubate transformation plates at 2-10 µE for 1-2 days.
      7. Increase the light intensity to 25-50 µE for 8-12 days.
        • This should yield >2 mm colonies that can be used for selection.

Alkaline Lysis

Stock Recipes

  1. Solution I
    Ingredient Final Solution Concentration
    Tris/HCl pH 8 25 mM
    EDTA 10 mM
    Glucose 50 mM
  2. Solution II
    Ingredient Final Solution Concentration
    NaOH 200 mM
    SDS 1%
  3. Solution III
    Ingredient Final Solution Concentration
    Potassium acetate 3 M

Protocol

  1. Inoculate 5 mL LB (with antibiotic for selection) with a single colony at 37°C, shaking at 360 rpm, overnight.
  2. Pellet 1.5 ml of the culture.
  3. Resuspend the pellet in cold solution I, vortex to ensure the pellet resuspends.
    • Note: DO NOT vortex after step 3.
  4. Add 200ul solution II, invert gently twice, place on ice for 3 minutes, to lyse the cells.
    • Note: DO NOT keep on ice for more than 5 minutes.
  5. Add 150 µl solution III, mix by inverting tubes gently, put on ice.
  6. Centrifuge in microfuge at full speed for 5 minutes, preferably at 3°C, to pellet cell membranes, etc.
  7. Transfer supernatant to new tube, add 400 µl pheno/chloroform to supernatant, mix by inverting, then spin in microcentrifuge for 5 minutes, max speed, to separate the layers.
  8. Transfer the upper phase to a new eppendorf add 600 µl 2-propanol, then leave on ice 10 minutes.
  9. Spin at room temperature, 5-10 minutes, max speed, to pellet DNA and RNA. Dump the supernatant, rinse pellet in 200 µl ice cold 70% ethanol.
  10. Centrifuge full speed 5 minutes, dump the supernatant, then leave tube open for 5 minutes to dry.
  11. Add 50 µL of tris/HCl, 10 mM, pH 8 to resuspend pellet.

Competent Cell Preparation

This protocol was provided by the iGEM advisors at the University of York with the advice that it is important that everything is kept ice cold and sterile.

  1. Inoculate 2 separate 250 mL flasks containing 50 mL of media with 500 µL of culture from an overnight.
  2. Grow with aeration until OD 650 reaches 0.5-0.6 (approx. 3 hours).
  3. Transfer the culture into sterile 50 mL Falcon tubes and chill on ice for 5 minutes.
  4. Spin down at 5000 rpm for 10 minutes at 4°C.
  5. Discard supernatant and resuspend the pellet in 10 mL ice cold sterile 0.1 M CaCl2
  6. Chill on ice for 20 minutes.
  7. Spin down at 5000 rpm for 10 minutes at 4°C.
  8. Discard supernatant and resuspend in 1.4 mL of 0.1 M CaCl2 mixed with 0.6 mL of 50% glycerol (ice cold).
  9. Aliquot 100 µL of culture into 1.5 mL sterile Eppendorfs.
  10. Freeze at -80°C.

E. coli Single Tube Transformation

This protocol was adapted from a protocol provided by iGEM.

  1. Materials
    1. DNA to be transformed
    2. 10 pg/µL Positive transformation control DNA
    3. Competent Cells (50 µL per sample)
    4. 1.5 mL Microtubes
    5. SOC Media (950 µL per sample)
    6. Petri plates with LB agar and antibiotic for selection
    7. Ice & ice bucket
    8. 42°C water bath
    9. 37°C incubator
    10. Sterile spreader or glass beads
    11. Pipettes and tips
    12. Microcentrifuge
  2. Procedure
    1. Label and pre-chill 1.5 mL tube.
    2. Thaw competent cells on ice (10-15 minutes)
    3. Pipette 50 µL of competent cells into 1.5 mL tube.
    4. Pipette 1 µL of resuspended DNA into 1.5 mL tube:
      1. Gently mix by pipetting.
      2. Keep all tubes on ice.
    5. Close 1.5 mL tubes, incubate on ice for 30 minutes:
      1. Gently agitate by flicking but return to ice immediately.
    6. Heat shock tubes at 42°C for 45 seconds.
    7. Incubate on ice for 5 minutes.
    8. Pipette 950 µl SOC media to each transformation:
      1. SOC media should be warmed to room temperature before use.
    9. Incubate at 37°C for 1 hour, shaking at 200-300 rpm.
    10. Pipette 100 µL of each transformation onto petri plates - spread immediately.
    11. Incubate transformations overnight (14-18 hours) at 37°C.
    12. Pick and screen for transformed E. coli.

Miscellaneous

List of Manufacturers' Protocols

  1. DNA extraction
  2. Qualitative PCR
  3. Cloning PCR
  4. PCR cleanup and gel extractions
  5. DNA assembly
  6. Enzymes for DNA analysis and manipulation

Fluorescence Microscopy

  1. Scrape off a small portion of a colony from the transformation plate.
  2. Use a sterile toothpick to streak cells on an object slide.
  3. Add a small drop of water and cover with a cover slip.
  4. Put the slide in the holder with the cover slip facing downwards.
    • Note the switch on the microscope that controls the backlight. This should be turned off to see fluorescence.
  5. Start at low magnification and increase, focusing on the smaller groups of cells.
  6. Once a target is selected, put immersion oil on the 100x objective lens and put it in place.
  7. The image can be diverted from eyepiece to screen if required.
    • Images can then be captured and saved.
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