Team:Newcastle/Notebook/Lab/Protocols/

Protocols

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: 30ml of PDMS makes 3 devices.

Materials

  • 50 ml Falcon tube. 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

Steps

  1. Pour 30ml of PDMS into a Falcon tube.
  2. Pipette 3ml of curing agent into the bottom of the tube containing the PDMS.
  3. Mix the PDMS and curing agent thoroughly by moving the pipette tip up and down.
  4. Place the falcon tube in a beaker and into a vacuum chamber. Ensure the falcon tube label is facing away from you so that you can more easily observe the liquid. Similarly have the vacuum chamber 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 PBMS to bubble up to the top of the tube.

    3. Turn off the vacuum pump and release the pressure, taking care 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 cure. Once cured the PDMS can be left with a lid on in the fridge and then used for several days.

  8. Prepare the mould.

    For our devices we used metal moulds consisting of a single chamber and two inlet channels for adding fluid.

    Figure 1: Our microfluidics mould with PDMS poured in.

    You will need to surround the sides of the mould with tape upto your desired height, ensuring that the mould is completely covered so that no PDMS leaks out.

  9. Cure the PDMS using a hot plate. Before using the hot place ensure that it is flat using a spirit level, adjust if necessary.

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

    2. Pop any remaining bubbles using a toothpick.

    3. Place on hotplate at 90C.

    4. Leave for 2 hours or overnight.

  10. Leave to cool and carefully peel the cured PDMS block from the mould.

  11. 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 endure 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 300ml flask.

    1. Add 0.5g of agarose.
    2. Add 50ml of 0.5x TBE.
    3. Mix, shaking by hand is OK.

    4. 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.

    5. Add 5µL of gel red using 0.5-5µL piptte.

  2. 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.
  3. 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 5µL 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.

    The gel may tear when you move it or remove the comb. If this happens check your gel concentration as you might need a higher value.

  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 90V.
  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 200µl 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.

Miniprep

  1. Harvest. Centrifuge 1–5 mL of the overnight LB-culture. (Use 1–2 × 109 E. coli cells for each sample.) Remove all medium.
  2. Resuspend. Add 250μL Resuspension Buffer (R3) with RNase A to the cell pellet and resuspend the pellet until it is homogeneous.
  3. Lyse. Add 250μL 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 350μL 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 12,000 × g for 1 minute. Discard the flow-through and place the column back into the wash tube.
  6. Add 500μL Wash Buffer (W10) with ethanol to the column. Incubate the column for 1 minute at room temperature. Centrifuge the column at 12,000 × g for 1 minute. Discard the flow through and place column back into the wash tube.
  7. Wash and remove ethanol. Add 700μL Wash Buffer (W9) with ethanol to the column. Centrifuge the column at 12,000 × g for 1 minute. Discard the flow through and place the column into the wash tube. Centrifuge the column at 12,000 × g 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 75μL 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 12,000 × g 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, 260/230 ratios.

M9 Minimal Media Preparation

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

TBE

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

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

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

TAE

As TBE but with the following ingredients:

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

LB

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

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

University of Reading's Microbial Fuel Cell Protocol

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 400mL of distilled or deionised water.
  2. Make up to 500mL 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 68.05g of KH2PO4 (potassium dihydrogen phosphate ) in 400mL of distilled or deionised water.
  2. Make up to 500mL 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 61.5mL of 1M K2HPO4 stock solution with 38.5mL 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 3.39g of potassium hexacyanoferrate (III) in 500mL 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 1.87g of methylene blue in 500mL of potassium phosphate buffer (see recipe above).
  2. Store in a labelled glass bottle at room temperature until required.

Glucose solution, 1M

  1. Dissolve 9g of glucose in 50mL 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 2.5g of dried yeast to 5mL of buffer solution and stir to produce a thick slurry.
  2. Next add 5mL 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 fbre 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 ft 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 5mL 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 ~10mL 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 0.4–0.6V and 3–50mA. 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