On this page you will find all of the protocols we've used throughout our project e.g. cloning protocols. Where any changes to these protocols have been made they are noted on the individual experiment pages in .

Protocol Library

Microfluidics Preparation Protocol

Note: 30 ml of PDMS makes 3 devices.


  • 50 ml Falcon tube or similar container. Note that the larger the tube the easier it is to pour.

  • PDMS polymer - Sylgard elastomer 184

  • PDMS curing agent. Caution the curing agent is photosensitive, keep covered.

  • Stopwatch

  • Sterile toothpick


  1. Pour 30 ml of PDMS into a Falcon tube.

  2. Pipette 3 ml of curing agent into the bottom of the tube containing the PDMS.

  3. Mix the PDMS and curing agent by moving the pipette tip up and down.

  4. Place the falcon tube in a beaker to keep it upright and return the pipette tip to its container. Ensure the falcon tube label is facing away from you so that you can more easily observe the liquid. Similarly have the vacuum jar pressure release valve towards you.

  5. Place the beaker containing the falcon tubes into the vacuum jar.

  6. Repeat the following steps until the PDMS stops bubbling and the gas is removed (approx. 4-5 times).

    1. Turn on the vacuum pump at 25-30 psi.

    2. Allow the PDMS to bubble up to the top of the tube.

    3. Turn off the vacuum pump and release the pressure. Be careful not to release the pressure too fast otherwise the beaker will fall over.

  7. Bring the PDMS up to vacuum using the pump and then leave for half an hour to degas. Once degassed the PDMS can be left with a lid on in the fridge and then used for several days.

  8. Set the PDMS using a hot plate. Before using the hot place ensure that it is flat using a spirit level. If necessary use the scrap PDMS wedges to flatten the plates.

    1. Pour the PDMS into the moulds ensuring an even covering.

    2. Pop any remaining bubbles using a toothpick.

    3. Place on hotplate at 100 °C.

    4. Leave for 2 hours or overnight.

  9. Leave to cool and carefully peel back the cured PDMS from the mould.

  10. Bond the device to a microscope slide using a plasma cleaner to activate the surface of the glass and the PDMS. Bake the chip for a further 30 minutes at 80°C to ensure good bonding.

Gel Electrophoresis Protocol

Preparing the Gel

Note: This protocol was written to produce a 1% agarose gel. Adapt the agarose and TBE concentrations as required to obtain concentration given in the lab book.

  1. Make up gel solution in 300 ml flask.
  2. Add 0. 5g of agarose.
  3. Add 50 ml of 0.5x TBE.
  4. Mix, shaking by hand is OK.
  5. Heat in microwave for 30 seconds, swill and then heat for a further 30 seconds. You may need to repeat the swilling and heating until the solution becomes molten
  6. Add 5 µL of gel red using 0.5-5 µL piptte.
  7. Allow the gel mix to cool whilst you prepare the chamber as follows.
    1. Check the gel container is not dirty, if it is clean it with water and then rinse with deionised water. Dry with blue roll. Do not use ethanol to clean the container as it may crack.
    2. Ensure gel tray is rotated 90 degrees and in the molding position.
  8. Once the mix has cooled (can be held for 30 seconds on your palm) it can be poured. If necessary you can cool the mix by running it under the tap whilst turning it. When you pour the gel ensure that the surface is flat and that if any bubbles appear you move them to the side using a pipette.
    1. Add your desired comb to the gel.
    2. Leave to set. Whilst the gel is setting you should remove your DNA from the freezer and allow it to thaw on ice. The instructions for running a gel can be found in a later section.

Running the Gel

  1. Remove DNA from freezer and place on ice to thaw.
  2. In a separate tube mix 5 µL of sample and 1 µL of loading dye.
  3. Once the gel has set rotate the gel 90 degrees so that the comb is nearest the negative electrode. This is because DNA is negatively charged and will migrate toward the positive electrode (red).
  4. Remove the comb from the gel.

  5. Load your DNA ladder into the first row of the gel.

  6. Load you DNA sample(s) into the subsequent rows of the gel.
  7. Run the gel for 40 minutes at constant voltage of 90 V.
  8. Image the gel using a UV plate, and check bands against ladder.

Transformation Protocol for E. cloni 10G & 10GF Cells

  1. Prepare nutrient agar plates (LB-Lennox) with antibiotic for selection. Ensure that Recovery Medium is readily available at room temperature.
  2. Chill sterile culture tubes on ice (17 mm x 100 mm tubes, one tube for each transformation reaction).
  3. Remove E. cloni cells from the -80 °C freezer and thaw completely on wet ice (5-15 minutes).
  4. Add 40 µL of E. cloni cells to the chilled culture tube.
  5. Add 1-4 µL of DNA sample to the 40 µL of cells. Stir briefly with a pipet tip; do not pipet up and down to mix, which can introduce air bubbles and warm the cells. For the pUC19 control, add 1 µL (10 pg) of DNA to another chilled culture tube containing 40 µL of cells.
  6. Incubate the cell/DNA mixture on ice for 30 minutes.
  7. Heat shock cells by placing the culture tubes in a 42 °C water bath for 45 seconds. Performing the heat shock in the 1.7 mL tube in which the cells are provided will significantly reduce the transformation efficiency.
  8. Return the culture tubes to ice for 2 minutes.
  9. Add 960 µL of room temperature Recovery Medium to the cells in the culture tube. When using these cells with a cloning kit, follow the Recovery Medium volume given in that kit manual.
  10. Place the tubes in a shaking incubator at 250 rpm for 1 hour at 37 °C.
  11. Plate up to 200 µL of the transformation on LB-Lennox or plates containing the appropriate antibiotic (Chloramphenicol).
  12. Incubate the plates overnight at 37 °C.
  13. Transformed clones can be further grown in any rich culture medium (e.g. LB).

Storage of Transformed Cells

  1. Grow overnight liquid cultures
  2. Add 200 µl of 100% glycerol to 800 µl of the overnight culture
  3. Store at -80 °C
  4. To recover the bacteria from the stock, open the tube and use a sterile loop to scrape off the frozen bacteria off of the top and streak onto an LB agar plate.
  5. Grow the plate overnight.

Plasmid Miniprep

  1. Harvest. Centrifuge 5 mL of the overnight LB-culture. Remove all medium.
  2. Resuspend the pellet. Add 250 µL Resuspension Buffer (R3) with RNase A to the cell pellet and resuspend the pellet until it is homogeneous.
  3. Lyse. Add 250 µL Lysis Buffer (L7). Mix gently by inverting the capped tube until the mixture is homogeneous. Do not vortex. Incubate the tube at room temperature for 5 minutes.
  4. Precipitate. Add 350 µL Precipitation Buffer (N4). Mix immediately by inverting the tube, or for large pellets, vigorously shaking the tube, until the mixture is homogeneous. Do not vortex. Centrifuge the lysate at >12,000 g for 10 minutes.
  5. Bind. Load the supernatant from step 4 onto a spin column in a 2 mL wash tube. Centrifuge the column at 12,000g for 1 minute. Discard the flow-through and place the column back into the wash tube.
  6. Add 500 µL Wash Buffer (W10) with ethanol to the column. Incubate the column for 1 minute at room temperature. Centrifuge the column at 12,000g for 1 minute. Discard the flow through and place column back into the wash tube.
  7. Wash and remove ethanol. Add 700 µL Wash Buffer (W9) with ethanol to the column. Centrifuge the column at 12,000g for 1 minute. Discard the flow through and place the column into the wash tube. Centrifuge the column at 12,000g for 1 minute. Discard the wash tube with the flow-through.
  8. Elute. Place the Spin Column in a clean 1.5 mL recovery tube. Add 75 µL of preheated TE Buffer (TE) to the centre of the column. Incubate the column for 1 minute at room temperature.
  9. Recover. Centrifuge the column at 12,000g for 2 minutes. The recovery tube contains the purified plasmid DNA. Discard the column. Store plasmid DNA at 4 °C (short-term) or store the DNA in aliquots at 20 °C (long-term).
  10. Test the purity by using the NanoDrop 260/280 nm, 260/230 nm ratios.

M9 Minimal Growth Media Preparation

  1. Make 1x M9 Salts by aliquoting 800 ml H2O and adding
    • 64 g Na2HPO4-7H2O
    • 15 g KH2PO4
    • 2.5 g NaCl
    • 5.0 g NH4Cl
    • Stir until dissolved
    • Adjust to 1000 ml with distilled H2O
    • Sterilize by autoclaving
  2. Measure ~700 ml of distilled H2O (sterile)
  3. Add 200 ml of M9 salts
  4. Add 2 ml of 1M MgSO4 (sterile)
  5. Add 20 ml of 20% glucose (or other carbon source)
  6. Add 100 µl of 1M CaCl2 (sterile)
  7. Adjust to 1000 ml with distilled H2O

TBE Gel Running Buffer

To make 1 L of TBE at 10x stock concentration add the following items to 600 ml of deionized water:

    • 108 g Tris base (FW = 121)
    • 55g boric acid (FW = 61.8)
    • 40 ml 0.5M EDTA (pH 8.0)

    Make up to a final volume of 1L using deionized water. To prepare a 1x working solution dilute 10× stock buffer with DNAse free deionized water in a ratio of 1 to 9.

TAE Gel Running Buffer

As TBE but with the following ingredients:

  • 242 g Tris base (FW = 121)
  • 57.1 ml glacial acetic acid
  • 100 ml 0.5M EDTA (pH 8.0)

LB Broth Growth Medium

  1. Add the following to 800 ml of deionized water
    • 10 g Bacto-tryptone.
    • 5 g yeast extract.
    • 10 g NaCl.
  2. Adjust pH to 7.5 with NaOH.
  3. Adjust to 1 L total volume using deionized water.
  4. Sterilize by autoclaving.

Alternatively a premixed LB powder can be used like Sigma EZMIX.

Microbial Fuel Cell Protocol (Based on the protocol of University of Reading

Protocol provided by Dr Ed Milner, Dr Paniz Izadi and Professor Ian Head. The original Protocol can be found here

Stock solutions

Potassium hydrogen phosphate 1M stock solution

  1. Dissolve 87.09 g of K2HPO4 (potassium hydrogen phosphate) in 400 ml of distilled or deionised water.
  2. Make up to 500 ml with distilled or deionised water.
  3. Store in a labelled glass bottle in a fridge at 3–5 °C until required.

Potassium dihydrogen phosphate, 1M stock solution

  1. Dissolve 68.05 g of KH2PO4 (potassium dihydrogen phosphate ) in 400ml of distilled or deionised water.
  2. Make up to 500 ml with distilled or deionised water.
  3. Store in a labelled glass bottle in a fridge at 3–5 °C until required.

Preparing the Phosphate Buffer

  1. Mix 61.5 ml of 1M K2HPO4 stock solution with 38.5 ml of 1M KH2PO4 stock solution.
  2. Make up to 1 litre with distilled or deionised water. This buffer should be used to make up all of the solutions required for the fuel cell.

Preparing the solutions

Potassium hexacyanoferrate (III), 0.02M

  1. Dissolve 3.39 g of potassium hexacyanoferrate (III) in 500 ml of potassium phosphate buffer (see recipe above).
  2. Store in a labelled glass bottle, wrapped in aluminium foil to exclude light, at room temperature until required. The solution should be used within six months of preparation.

Methylene blue, 10mM

  1. Dissolve 1.87 g of methylene blue in 500 ml of potassium phosphate buffer (see recipe above).
  2. Store in a labelled glass bottle at room temperature until required.

Glucose solution, 1M

  1. Dissolve 9 g of glucose in 50 ml of potassium phosphate buffer (see recipe above).
  2. Use immediately or within 24 hours of preparation as the solution is not sterile and will support the growth of contaminating microorganisms.

Preparing the Fuel Cell Parts

Each compartment of the fuel cell is made of two Perspex® parts. Four neoprene gaskets are provided that can be sandwiched between the parts to prevent leaks from the cell. If desired, however, the two parts that make up each compartment can be stuck together using clear silicone sealant, so that only two gaskets will then be needed on each side of the cation exchange membrane.

Assembling the Fuel Cell Parts

  1. Before you start to assemble the fuel cell, rehydrate the dried yeast in potassium phosphate buffer, pH 7.0. To do this, add 2.5 g of dried yeast to 5 ml of buffer solution and stir to produce a thick slurry.
  2. Next add 5 ml of 1M glucose solution to the yeast slurry and stir well to mix. Don’t be tempted to rehydrate the yeast in the glucose solution or to mix the glucose solution with the phosphate buffer before adding the yeast — if you do this, the yeast will not rehydrate as readily.
  3. Put the yeast suspension to one side, then cut out and fold two carbon fibre electrodes as shown in the picture on the right.
  4. Insert one electrode into each chamber of the fuel cell. Assemble the two halves identically, so that when they are held together, there will be room to insert a syringe into one of the small holes over each chamber.
  5. Cut out two pieces of J-Cloth® or similar non-woven fabric — one to fit into each chamber of the fuel cell. Place one in each chamber on top of the electrodes. The purpose of the J-Cloth® is simply to prevent the electrodes from touching the cation exchange membrane.
  6. Place a neoprene gasket on each half of the fuel cell, then place the two halves together with the cation exchange membrane sandwiched between them.
  7. Pass the four bolts through the holes in the outer parts of the cell and tighten the wing nuts. Do not over-tighten the nuts, as this may distort the cell and allow liquid to weep out.
  8. Stand the assembled fuel cell on a Petri dish base or lid to catch any liquid that leaks from the cell.
  9. Add 5 ml of 10mM methylene blue solution to the yeast suspension. Stir well, then syringe the mixture into one chamber of the fuel cell.
  10. Use a clean syringe to add ~10 ml of 0.02M potassium hexacyanoferrate (III) solution to the other chamber of the fuel cell.
  11. Connect a voltmeter or multimeter to the electrode terminals using crocodile clips. Fuel cells of this type typically generate 0.4–0.6 V and 3–50 mA. A current should be produced immediately — if the meter registers zero, check the connections and ensure that the carbon fibre electrodes are not touching the cation exchange membrane