Difference between revisions of "Team:Kent/Experiments"

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Experiments & Protocols

Basic Protocols

Production of Lysogeny broth (LB)

For 1 litre of LB a mixture of 10g of sodium chloride, 10g peptone, 5g of yeast extract as well as 1 litre of distilled water in a glass bottle. We then used a magnetic spinner to help mix the powders with the water, we avoided shaking the glass bottle as it would cause froth and waste some of the LB.
When making the LB we also made another litre batch and added 15g of agar extract to be able to grow bacteria on plates.

Production of SOB medium and magnesium stock

Bringing together 20g of tryptone, 5g of yeast extract, 0.584g of NaCl, 0.186g of KCl and mixing it with 990ml of millipure water (using the magnetic mixer again) which was then put in to autoclave to sterilise it, after it was taken out and let for it to cool down to below 60 o C.
10ml of 2M Mg 2+ stock was then added and then brought to 100ml with millipure water, 0.2mm filter sterilize was then used

Production of SOC medium and glucose stock

Once again bring 20g of tryptone, 5g of yeast of extract, 0.584g of NaCl, 0.186g of KCL, and then bring 970 ml with millipure water and use the magnetic mixer once again, this was also then put in to autoclave.
10ml of 2M Mg 2+ stock and then bring it to 100ml with milllipure water, filter sterilize it with 0.2m and then final add 20ml of 1M glucose stock.

Production of Glycerol stock

If you wish to store bacteria long term, you will need to create a Glycerol Stock after inoculating an overnight liquid culture

Once bacterial growth has been achieved, 500μL of the overnight liquid culture needs to be added to 500μL of 50% glycerol in a 2mL tube where it should be gently mixed
The glycerol stock should then be frozen at -80 o C
- Successive freeze and thaw cycles will reduce the stocks shelf life

Running Agarose Gel

After the cells have been miniprepped and the plasmid put through a restriction digest, the agarose gel can be run.

Make up some agarose. This is done by taking 0.5g of agarose powder and putting it in a 250ml sterile conical flask, with 50ml of TAE buffer, then microwaving it in small pulses (20 seconds then swirling it around) until it is dissolved. Don’t overheat it or it will evaporate too much. Make up the evaporated volume to 50ml with distilled water.
Add 1 vial of cybersafe (ask technical services for a tube of it and add all of it)
Line the white sides of the tank with the agarose solution, to seal it and prevent leakage. Use a p1000 pipette set to 1ml. Let it dry (about 5 mins max)
Then pour all the agarose/sybrsafe solution into the tank and put in the comb. Let it set and solidify (maximum 30 mins)
When the gel has set, remove the comb from the tank (gently!) and then cover the whole tank with TAE buffer, so there’s at least half a centimetre of TAE covering the gel.
Now, the samples need to be loaded. Load some DNA markers (ask technical services for a tube of this and load the whole tube) into well 1( left hand side) and then choose what you load into wells 2, 3, and 4 etc. (make sure you note what’s in each lane!)
Load all of your digests into the wells 2,3, and 4.
Plug into a power supply and put the cover on. Run for 40 mins to an hour at 80v. The amps don’t matter.
Once the visible markers have reached the half way point of the tank, turn off the power supply and drain the TAE buffer form the tank. Remove the gel with a spatula and place in a UV imaging box. Take an image of the gel under UV light, save it onto a USB stick.

Overnights protocol

After a transformation has been run and plates have been streaked and patched, overnight cultures will need to be made:

Add 10mL of LB broth (not agar) into as many autoclaved conical flasks as needed
Add 10uL of Chloramphenicol into each conical flask as well
Using a pipette tip, scrape up some of the cell colonies on the agar plates prepared beforehand and drop it into the conical flask
Cover up the flask using aluminum foil
Incubate the cultures at 37oC and 180 rpm

Protocol for transformation/ heat shock

This requires chemically competent cells to be made beforehand. These must be stored at -80 degrees Celsius:

First, the competent cells and the plasmid intended for transformation must be thawed on ice. Additionally, some 1.5ml Eppendorf tubes should be chilled on ice, along with some pipette tips.
100ul of the chemically competent cells are then pipetted (using the chilled pipette tip) into a chilled Eppendorf tube.
5ul of DNA is the pipetted into the tube with a chilled tip. This tube is then stored on ice for 30 minutes.
The tube is then placed in a water bath at 42 degrees Celsius for precisely 90 seconds. After 90 seconds is up, the tube is transferred back to ice for 2 minutes.
900ul SOC medium is then added into the tube (with a normal pipette tip, doesn’t need to be chilled) and then mix very gently by pipetting up and down inside the tube.
The tube is then incubated at 37 degrees Celsius for 45 minutes. The tube should not be shaken at all at this point.
100ul of the transformation mix is then pipetted into the centre of a plate containing LB agar and the appropriate antibiotic (for example, for plasmid pSB1C3, Chlorophenicol should be used, and for pSB1A3, use Ampicillin). Use 1ul of antibiotic for each ml of agar.
In sterile conditions (Bunsen burner, gloves cleaned with IMS (ALLOWED TO DRY)), spread the bacteria around the plate by keeping the lid as closed as possible and inserting the spreader then turning the plate around to spread the cells. Then immediately close and store upside down.
The remained of the transformation mix is then spun at the highest possible speed for 2 minutes. The resulting pellet is then resuspended in 100ul of the existing medium and plated onto the LB and antibiotic plate.
Incubate in a 37-degree Celsius incubator for 16-18 hours.
Any colonies that result from this should be plated on a patch plate.
This is done by taking a plate of LB agar with the appropriate antibiotic and dividing it up into sections by drawing a grid on the bottom. These sections are numbered and then using a sterile pipette tip, the colonies are gently streaked in each section- 1 colony per section.

Transformation Guidelines (QuickChange Protocol)

Store all XL1-Blue supercompetent cells at -80 o C (prevents loss of efficiency) as they are sensitive to the smallest of temperature variations, even transferring tubes from one freezer to another will result in loss of efficiency
Storage Conditions:

XL1-Blue supercompetent cells should be stored at the bottom of a -80 o C freezer
They should be placed at -80 o C directly from dry ice shipping container

Keep the XL1-Blue supercompetent cells on ice at all times
Essential that BD-Falcon polypropylene tubes are places on ice before cells are thawed
Cells must be aliquoted directly into prechilled tubes (Use of 14-mL BD Falcon Polypropylene Round-Bottom Tubes)
These tubes must be used (BD Biosciences Catalog #352059) for the transformation protocol
The heat-pulse steps’ duration is critical and has been optimized for the thickness as well as shape of these tubes

Optimal efficiencies observed when cells are heat pulsed for 45 seconds
Heat pulsing for at least 45 seconds is recommended, allowing for slight variations in incubation length
Efficiencies noted to decrease sharply when pulsed for <30 seconds or >45 seconds
This defined window of highest efficiency for the XL1-Blue cells results from heat pulse in step 3 of transformation protocol

To the LB agar, add
- 80 µg/ml of 5-bromo- 4-chloro- 3-indolyl- β-D-galactopyranoside (X-gal)
- 20 mM isopropy-1- thio β-D galactopyranoside (IPTG)
- Appropriate antibiotic
These are all added to prepare the LB agar plates for blue-white colour screening
Alternatively
- 100 μl of 10 mM IPTG and 100 μl of 2% X-gal can be spread on LB agar plates 30 minutes prior to plating transformations
The IPTG must be prepared in sterile dH2O
The X-gal must be prepared in dimethylformamide (DMF)
IPTG and X-gal MUST NOT be mixed before being pipetted onto the plates since the chemicals may precipitate.

Interlab Protocols

Calibration of OD 600 Reference Point

LUDOX-S40 must be used as a single point reference with the aim to attain a ratiometric conversion factor, which in turn will be used to transform absorbance data into standard OD 600 measurement.
Standard 1 cm path length spectrophotometer readings are instrument dependent, while plate readers possess a path length less than 1 cm and are volume dependent.
Therefore, in this situation, there are 2 key objectives:

Ratiometric conversion to transform Abs 600 measurements into OD 600 measurements
Accounting for instrument differences

Before starting the protocol, path length correction must be switched off. This is because scattering increases with longer wavelengths therefore adjustment is confounded by scattering solutions such as dense cells. However, many plate readers have automatic path length correction which is based on volume adjustment that uses ratio of absorbance measurements at 900 + 950 nm. LUDOX solution is only weakly scattering so will produce low absorbance values
Use same cuvettes, plates and volumes that are going to be used in cell based assays
Materials:

1 mL 100% LUDOX
H 2 O
96 well plate (black with flat, transparent/clear bottom)

Method

100 µl of LUDOX should be added into wells A1, B1, C1 and D2 (or 1mL into cuvette)
100 µl of H 2 O should be added into wells A2, B2, C2 and D2
Measure absorbance 600nm of all samples in all standard measurement modes in instrument
Record data

Production of Fluorescein stock solution

Spin down the Fluorescein stock tube and ensure the pellet is at the tubes' bottom
Prepare 2x fluorescein stock solution (100 µM)
- Resuspend Fluorescein in 1mL 1xPBS
- Ensure Fluorescein is properly dissolved
  After the resuspension, pipette up and down and examine the solution in the tip (if particulates are visisble, continue to mix solution until they disappear)

Dilute the 2x Fluorescein stock solution
- With 1xPBS to make 1x fluorescein solution
- With resulting concentration of fluorescein stock solution 50 µM (500 µL of 2x fluorescein in 500 µL 1x PBS to make 1 mL of 50 µM (1x) fluorescein solution)

Fluorescein Fluorescence Standard Curve

A dilution series of Fluorescein in 4 replicates must be prepared where the fluorescence is measured in a 96 well plate in standard mode on a plate reader. A standard curve will be generated of fluorescence of fluorescein concentration. This will be used to correct cell based readings to an equivalent fluorescein concentration, which will then be converted into a GFP concentration.

Materials

Fluorescein
10mL 1xPBS (Phosphate Buffered Saline)
96 well plate (black with flat, transparent/clear bottom)

Method
Serial dilutions need to be performed across columns 1-11 Column 12 must contain PBS buffer only
The plate will initially be setup fluorescein stock in column 1 and equal volume of1xPBS in columns 2-12

Add 100 µL of PBS into wells A2-A12, B2-B12, C2-C12 and D2-D12
Add 200 µL of Fluorescein 1x stock solution into A1, B1, C1 and D1
Transfer 100 µL of Fluorescein stock solution from A1 into A2
Mix A2 by pipetting up and down 3x and transfer 100 µL into A3 Repeat the process for A3 into A4, A4 into A5, etc. until A11
Mix A11 by pipetting up and down 3x and transfer 100 µL into liquid waste
Repeat dilution series for rows B, C and D
Measure fluorescence of all samples in all standard measurement modes in instrument
Record the data

Measurement notes

The plates can now be measured in the plate reader
Standard GFP settings must be used (same as those used when measuring the cells):
- Excitation 485nm
- Emission 530/30
- Turn off path length correction
Would be ideal to repeat measurements with different settings
- Generates series of standard curves to choose from
Use number of settings that affect sensitivity (gain and/or slit width)
- Also consider orbital averaging, top/bottom optics

Cell Measurement Protocol

The calibration measurements should be performed before the measurements on the cells are performed. This allows that the measurement process is understood and that the cell measurements are taken under the same conditions.
Materials

Competent cells (E.coli strain DH5-alpha)
LB (Luria Bertani) media
Chloramphenicol (stock concentration 25 mg/mL dissolved in EtOH – working stock 25 ug/mL)
50 mL Falcon tube (covered in foil to block light)
Incubator at 37oC
1.5mL Eppendorf tubes for sample storage
Ice bucket
Pipettes
96 well plate (black with flat, transparent/clear bottom)

Calcium Chloride Competent Cells

Prior Preparation

Autoclave 50mM Calcium Chloride and keep it cold at about 4 o C
For the starter cultures
- Add a colony of E.coli DH5cells to 5mL of LB
- Incubate at 37 o C overnight

Keep cells on ice at all times where possible
To 100mLs of LB, add 100uL of cells from the overnight culture
Let it grow at 37 o C and 250 rpm (until it reaches OD 600 ~0.6-0.8)
Place cells on ice immediately to cool them once the correct OD 600 has been reached
Centrifuge at max speed for 10 mins and 4 o C
Discard supernatant
Resuspend the pellet in 50% of the original volume with ice-cold 50mM CaCl 2; In a 5omL culture, add 25mL CaCl 2
Allow them to sit on ice for 30 mins
Centrifuge at max speed for 10 mins at 4 o C
Discard the supernatant

Cell Line
Sterile LB
10mM sterile and chilled Calcium Chloride
Dry ice
Acetone

Inoculate the cells (either 1:50 or 1:100) into 50mL of LB
Grow them at 37 o C until OD600 is around 0.4-0.5
Place on ice for 10 minutes while Falcon tubes are pre-chilled
The cells should be harvested at 3000 rpm, 4C for 8 minutes
The pellet then needs to be resuspended in 1mL of 100mM CaCl 2 and 30% (v/v) glycerol
The resulting solution needs to be aliquoted into chilled Eppendorf tubes (100uL per tube)
Place each Eppendorf tube into an acetone dry ice bath to snap freeze them
Then store at -80 o C

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