Difference between revisions of "Team:Oxford/Protocols"

1.1 Gel Electrophoresis

For a large 1% gel, prepare 200mL agarose

1. Heat 2g agarose in 200ml 0.5x TBE for 2 minutes under full power in the microwave (use a 500mL Duran bottle, and place a weighing boat underneath it to prevent the causing of a mess in the event the mixture boils over; DO NOT fully tighten the Duran cap).
2. Check if the agarose has been fully dissolved. Heat it further if gel strands are visible.
3. Hold the lid with paper and gently swirl.
4. Leave the agarose solution to cool at 50℃ for 20 minutes.
5. Pour agarose onto gel plate in a setting tray with appropriately-sized combs already fixed onto it, and leave for 20 minutes to let it set.
6. When the agarose has set, remove the combs and transfer the gel plate from the setting tray to the electrophoresis chamber.
7. Flood the gel plate with 0.5x TBE buffer up until right above the top of the wells.
8. The gel should be positioned such that the positive (red) electrode is on the far side of the gel from the wells, as the negatively-charged DNA will migrate towards the positive electrode.
9. Load 10\(\mu\)L DNA ladder in lane 1 and 20\(\mu\)L PCR product in subsequent lanes.
10. 120V for a big gel (200mL agarose) or 80V for a small gel(100mL agarose).

1.2 Restriction Digest PCR or Plasmid DNA

• Use enzymes and buffer according to "Master Table".
• Defrost and shake buffers.
• Keep enzymes in yellow freezing block and keep out of freezer for as short a time as possible.

A point to note concerning the volume of restriction enzyme

• 0.5\(\mu\)l for PCR DNA
• 1.0\(\mu\)l for Plasmid DNA
• However, if you are doing a test digest (i.e. after a mini-prep) use 0.5\(\mu\)l enzyme, despite digesting a plasmid

1. Incubate at 37℃ for 2 hours (ThermoMixer program 3) with shaking at 300rpm.
2. Heat inactivated for 30 minutes at 95℃
3. Dephosphorylate the plasmid using 1\(\mu\)l CIP at 37℃ for 3 minutes

1.22 Nanodrop

1. Clean stage with 1\(\mu\)l water and tissue.
2. Make a blank reading using 1\(\mu\)l of water and wipe off.
3. Make another blank reading using 1\(\mu\)l of elution buffer and wipe off.
4. Measure concentration of 1\(\mu\)l of each sample.

1.3 Ligation

Overnight protocol

Defrost T4 DNA Ligase on ice.

Keep in freezing block when on bench and add last to the reaction mixture.

'Mass of vector DNA : Mass of insert DNA' roughly 1:3.

Generally, because you only get 50\(\mu\)L from the plasmid digest, split evenly between how many inserts there are.

The component volumes are:

Incubate at 16℃ overnight.

1.4 Preparation of Competent Escherichia coli Cells

Overnight protocol

DH5ɑ is stored at -80℃

1. Defrost and inoculate in 5mL of LB in a 125 mL conical flask (volume of LB 10% of flask volume so as to achieve sufficient aeration).
2. Grow culture overnight at 37℃.

Next Day

Turn on the centrifuge and cool to 4℃.

Always keep TFBI and TFBII on ice.

Important to keep volumes accurate; else your cells will grow at different rates and OD. s will be all over the place.

1. Transfer 1ml of overnight culture into 100ml LB in a 500ml conical flask.
2. Incubate at 37℃ until OD_600nm reaches 0.4-0.6. It is good to stop at OD = 0.35, as the bacteria are now replicating exponentially i.e. will only take 20 more minutes until OD = 0.7 (which is far too high).
3. While incubating, pre-chill 20 eppendorfs and 2x50mL Falcon tubes (which are found in the cupboard next to the sink) to 4℃ in ice bucket.
4. Once OD is correct, decant culture into 2 x 50ml falcon tubes . Once this is done, the cells must never be higher than 4℃.
5. Centrifuge at 2000 rpm at 4℃ for 20 mins. Make sure you are at the centrifuge when this 20 minutes is up to rapidly proceed to the next step. Close the centrifuge lid to maintain 4℃.
6. Discard supernatant and resuspend pellets in 1ml TFBI.
7. Add further 10ml TFBI using 10ml electronic pipette.
8. Leave on ice for 20 mins (or more).
9. Centrifuge at 2000 rpm at 4℃ for 10 mins (shorter duration as cells are already now permeabilized).
10. Discard supernatant.
11. Gently resuspend pellet in 2ml TFBII i.e. don’t vortex, just flick pellets in the TFBII, which has glycerol for frost protection (prevent crystal formation which lyses cells).
12. Now we have total 4mL of cell-TFBII suspension. Aliquot ~200ul into ~20 pre-cooled eppendorf tubes.
13. Store at -80℃ in the freezer in the back room.

1.5 Transformation

Overnight Protocol

To transform cells you need to prepare agar plates. This can be done when you have a spare hour during the day. Plates can be left on the bench (lid side down) until you need them. Plates can also be made during the transformation protocol.

200ul aliquots of competent cells are stored at -80C in the back. These take around 30 minutes to defrost and so these need to be taken out of the freezer and put on ice to defrost. 100ul of competent cell = 1 transformation.

Preparing the cells for transformation

For ligated DNA:

1. Put the eppendorfs of ligated DNA into ice to cool to 4℃.
2. Draw 100\(\mu\)l of competent cells of each aliquot to each 50ul ligation product.
3. Incubate on ice for 30 minutes. If you are preparing plates as you go along, the LB agar should be done microwaving at this point.
4. Bring ice bucket to the water bath.
5. Heat shock - transfer straight from ice bucket into 45℃ water bath for 45 seconds.
6. Transfer back to ice for 1 minute.
7. Add 800\(\mu\)l LB broth to each tube and incubate at 37℃ gently shaking for 1 hour.

For plasmid DNA:

1. Thaw the plasmid DNA for transformation or use ligation product.
2. Pre-chill number of eppendorfs to match number of transformations.
3. Thaw appropriate number of 200\(\mu\)l aliquots of competent E. coli cells to prepare 100\(\mu\)l eppendorfs to match number of transformations.
4. Draw 100\(\mu\)l out of each aliquot (competent cells) and transfer into empty pre-chilled eppendorfs.
5. Transfer the full volume (usually ~50\(\mu\)l) of the plasmid DNA into the respective E. coli aliquots (we use all of it as we expect the plasmid concentration to be relatively low). For the stock pSB1C3, add only 1\(\mu\)L of the plasmid to the 100\(\mu\)L of cell suspension as the plasmid is at much higher concentration than for e.g. ligation products.
6. Incubate on ice for 30 minutes. If you are preparing plates as you go along, the LB agar should be done microwaving at this point.
7. Bring ice bucket to the water bath.
8. Heat shock - transfer straight from ice bucket into 45℃ water bath for 45 seconds.
9. Transfer back to ice for 1 minute.
10. Add 800\(\mu\)l LB broth to each tube and incubate at 37℃ gently shaking for 1 hour.

1.6 Growth and Culture of Bacteria

Overnight protocol

This process significantly increases the amount of plasmid that contains biobrick that we want. Plasmids can be extracted later.

Antibiotics are in the freezer - ampicillin needs defrosting before you start.

The LB you use has to be transparent (cloudy = contaminated).

Sterile Technique

The tip of each test tube and Duran bottle must be sterilised with a Bunsen burner each time when you are preparing the tubes and each time you transfer a colony.

[Arrange your hands in a way that you have your thumbs free and lids do not get placed on the desk. This is difficult to explain so ask someone who has done it before to show you.]

1. Choose three colonies from each plate. The colony should not be too small or too large and should be reasonably spaced from the others.
2. Label a test tube for each colony.
3. Pour 5ml LB broth that has antibiotic to be diluted 1/1000 fold to each tube i.e. if you have 10 plates and you are preparing 30 tubes, you will need 30x5 = 150ml LB with 150\(\mu\)l antibiotic already added. Therefore, each tube contains 5ml LB and 5ul antibiotic.
4. Using inoculation spatulas, pick colony and transfer to appropriate tube. When taking an inoculation spatula from the packet, be careful not to reach into the packet; instead, push handle out of packet to keep sterile.
5. Push down and bring the colony directly into LB without touching the sides.
6. Remove the test tubes from the rack and place in 37℃ overnight.

1.7 Mini-prep

E.Z.N.A. Plasmid DNA Mini Kit I

If you haven’t done a mini-prep before ask someone who has to go through it with you.

• Carry out all optional steps except equilibration step
• Repeat 1st centrifugation (step 2) until all the LB broth has been spun down and all the E. coli have been collected - will help increase yield later
• After centrifugation in step 2, pulse tubes before the excess supernatant was removed through pipetting
• After addition of solution I/RNase, vortexing/vigorous shaking of the tubes should be avoided to prevent shearing of nucleus and undesirable accidental extraction of chromosomal DNA
• After addition of solution I/RNAse, resuspension of pellet can be done by dragging the tube along an Eppendorf rack
• Steps 6 and 7 (involving solutions II and III need to be carried out in quick succession (adhering to the short incubation time) to ensure good results. It is advisable to do these two steps in pairs as in step 6 the tubes need to be tightly capped once solution II is added
• The inversion in step 6 needs to be done gently so that genomic DNA of the bacteria are not extracted along with the desired plasmid DNA
• After addition of solution II, the waiting time before proceeding to the next step should not be more than 5 minutes
• The inversion in step 6 needs to be done gently so that genomic DNA of the bacteria are not extracted along with the desired plasmid DNA
• The precipitate formed in following step 7 does not pellet well after centrifugation in step 8, and hence the supernatant needs to be removed immediately to prevent resuspension

• Elution step:

  • place elution buffer in 55℃ water bath (50\(\mu\)l per miniprep)
  • pipette warmed elution buffer onto the HiBind column and let it sit for 3 mins
  • centrifuge at max speed for 1 minute
  • pipette up filtrate and pipette back onto HiBind column
  • centrifuge at max speed again

→ Nanodrop → Restriction Digest → Gel electrophoresis → Sequence

3.1 SDS-PAGE

Running the gel

1. Prepare 1x SDS loading dye containing a final NaOH concentration of 50\(\mu\)M.
2. Resuspend the pellets using 30\(\mu\)L of said dye, and boil at 99℃ for 5 minutes or until the pellet dissolves to give a blue solution.
3. If the resulting solution is yellow, adding 5\(\mu\)L of 20 mM NaOH should turn it blue. Be careful when opening lids of hot eppendorfs - if the interior of the tube is steamy, let it cool down and subsequently spin it down to pull down the liquid first.
4. Obtain PAGE gel cassette from cold room, and dilute some 20x SDS buffer (found on top of bench) to 1x (800mL typically needed to fill PAGE tank). NB: Use the tank without the yellow strips on the inside.
5. PAGE tank can be found in one of the cabinets at Tom’s bench.
6. Load cassettes and 1x buffer into tank.
7. Obtain ladder (SDS 2-colour dye) from Jia’s freezer, load 15uL into well in cassette.
8. Load 15\(\mu\)L of each sample into wells.
9. Run gel for at least 40 minutes at 100mA per gel.
10. Once the dyefront is at a desirable position, remove gel from cassette and place it in a square petri dish. Stain with Instant Blue for 1 hour but preferably overnight.

3.2 Western Blot

Day 1

Follows on from SDS-PAGE

• In SDS-PAGE, must load 10 μl of benchmark ladder in lane 1 and 10 ul 2-Colour SDS Marker in the lane 2, so that the orientation of the gel can be determined and the efficiency of blotting checked later.
• You will need approx. 1L cold blotting buffer; to make 1L.
  • 3g Tris base
  • 14.4 g glycine
  • 200 ml methanol
  • 10 ml 10 % SDS
  • then make up to 1L with water

How to make a sandwich

Blot on to PVDF membrane using the following layout in your sandwich:

How to Make a Sandwich¹

• The black face of the blotting cassette is the 'negative' side whilst the white face is the 'positive' side
• Construct the sandwich layer by layer, from the black face of the casing upwards to white face
• Sandwich must be made whilst it is submerged in a Tupperware box of cold blotting buffer
• Use tweezers to handle the layers
• Every layer must have air bubbles removed using the plate spreading tool

Components

• 2 sponges
  • Found in cupboard under microwave
  • Soak in blotting buffer, smooth out bubbles, then place on cassette
• 6 (2 layers of 3) pieces of filter paper
  • Should be cut to the same size as the gel (8cm x 9cm)
  • It is found between the drawers under the nanodrop machine.
  • Soak 3 pieces of paper at once
• Gel from SDS-PAGE
  • Remove the gel from the plastic casing, place in blotting buffer and then onto the cassette
• PVDF Membrane
  • Found in the labelled drawer under the gel running area
  • The membrane is white and is protected within two sheets of blue paper
  • Try not to touch the membrane itself, even with gloves on
  • Instead handle membrane with tweezers
  • Before immersing in blotting buffer, soak in methanol (found under fume hood) for 5 minutes
  • Cut the edge of the PVDF membrane in the top left corner, ie nearest the well of lane 1 (This will allow you to keep track of which side of the membrane you have blotted on to later!)

After adding all components according to the diagram above, close the blotting cassette and place in the running tank.

• If only one gel is to be blotted then fill the other space with a second cassette containing two blotting pads.

Insert a cooling block containing frozen Milli-Q.

• These are found in the lowest drawer of the fridge next to the iGEM fridge

Fill tank with cold blotting buffer, place tank in an ice bucket and run at 0.5 A 1 hour.

Disassemble blotting apparatus and place blot in 5% milk powder in PBS with shaking overnight.

3.3 Ni²⁺ Affinity Chromatography

Day 2

First thing in the morning prepare the growth medium - the volume is dependant on how much you wish to purify - 20 mL total with 1 mL of overnight culture. Grow flasks at 37C with shaking at 225 rpm until OD₆₀₀ is 0.4 - 0.6.

Induce protein expression at appropriate temperature incubator with 0.2% ara for 4 hours.

Supernatant purification

• Spin cultures at full speed for 15 minutes
• Transfer the supernatant into labelled falcon tubes
• Filter supernatant

Lysate purification

• Keep on ice throughout
• Spin culture at full speed for 15 mins and discard the supernatant.
• Add 15 ml of resuspension buffer
• Vortex
• Add 20 mg lysozyme
• Sonicate - 4 min, 5 seconds on, 15 seconds off
• Filter

Setting Up Columns

Acquire the Ni-NTA Agarose slurry from the fridge and gently swirl until the Nickel beads are resuspended.

Pour 1 mL Ni-NTA Agarose slurry into a chromatography column. Allow to settle for at least 10 minutes.

Equilibrate the column with 30 mL resuspension buffer or Milli-Q.

Apply the filtered supernatant or lysate to the equilibrated nickel column.

Wash the column with wash buffer.

Elute the protein using the elution buffer. Collect 6 1 ml fractions of the eluate. Protein usually elutes in the first three/four fractions.

assay the fractions for protein using, for example, a Bradford assay.

Pool the fractions containing significant quantities of protein and dialyze overnight (against Water using a 10,000 MW cut-off dialysis membrane; these can be stored at -20C)

5.1 Fluorescence Assay

Perform media swap:

1. Spin down 1 mL stationary phase overnight culture
2. Discard supernatant
3. Resuspend in 1 mL unsupplemented M9
4. 1/100 dilution M9 media
5. Grow to mid-log phase which gives OD value of 0.2-0.5.
6. 100µL mid-log phase culture to each well
7. Run OD script how you would for toxicity; number of cycles depends on how much time you have but roughly 60-80 cycles
8. Also set to grow a fluorescence curve
9. Right click - for toxicity OD 600, put another protocol for GFP

8.1 Agaraose Bead Preparation

In a fume cupboard, break up a standard (polystyrene) petri dish into small pieces and dissolve in the minimum amount of ethyl acetate .

Prepare a required volume of 1.5 % Agarose solution in breaker with screw cap. Microwave on high for 2 minutes and then cool to 40 °C in water bath, when microwaving ensure the screw cap is placed on loosely.

Remove agarose solution from water bath and bring to laminar airflow cupboard. Pour the agarose into petri dish to a depth of 1cm. Allow to set, this should take roughly 15 minutes.

Still under laminar flow, use an autoclaved 1 cm diameter hole borer to core out the required number of identical agarose cylinders and place in a second petri dish and allow drying.

Bring uncoated ‘beads’ to fume cupboard. Using autoclaved needles, pick up individual beads and dip in ethyl acetate-polystyrene solution, stand each bead on needle upright in fume cupboard on blob of blu-tac to allow ethyl acetate evaporation and the coating to set. When the coating is almost set the beads should be able to be handled through gloves without damaging the coat. Remove needle and mould the polymer coat over the needle hole, sealing them.

8.2 Sodium Alginate Bead Prepartation

Make up required volume of 1.2% alginate solution in dH₂O. Slowly add Sodium Alginate to hot water and add a magnetic stirrer bar. Then leave to stir until all solid is dissolved.

Transfer this mixture in a 2mL syringe with a needle attached and drop the mixture into a solution of 0.1M calcium chloride. Vary the size of the syringe and height from which you drop and record the effect on the beads. Leave the beads in the CaCl₂ solution for 5 minutes and then filter the beads out.

8.3 Diffusion Rates

First make up solutions of Sodium Fluorescein of the following concentrations (µM) by serial dilution of 0.02M Sodium Fluorescein:

1, 2, 3, 4, 5, 10, 20, 30, 40, 50.

Then make up the following concentrations (%) of Crystal Violet by serial dilution of 0.1% Crystal Violet solution:

0.05, 0.01, 0.005, 0.001, 0.0005, 0.0001, 0.00005, 0.00001, 0.000005, 0.000001.

Make up 100mL of 1.2% Sodium Alginate solution and leave to cool.

Using the UV-VIS spectrometer measure the absorption of 1mL of Sodium Fluorescein solution at each concentration. The wavelength of light should be set to 480nm, as this is roughly the wavelength at which the absorption maxima lies. Then measure the absorption of 1mL of Crystal Violet solution. The wavelength of light should be set to 590nm, as this is roughly the wavelength at which the absorption maxima lies. This data can then be used to plot a calibration curve.

To make the beads, mix 2mL of 50µM Sodium Fluorescein solution with 18mL of Sodium Alginate, or mix 2mL of 0.1% Crystal Violet with 18mL of Sodium Alginate. Filter the beads and wash with Milli-Q.

Place the beads into 100mL (±5%) of Milli-Q.

Remove a 1mL portion of reaction solution out every 10 minutes and measure the absorption at 480nm for Sodium Fluorescein or 590nm for Crystal Violet.