- Determine the concentration of the DNA sample by running both the vector and insert ona 1% agarose gel and comparing the bands intensity with the ladder (concentration known).
- Calculate how much solution is needed to obtain desired total amount of DNA for digestion.
- The volume of DNA solution can be no more than 70% of the total solution. Therefore calculate the total volume of digestion (probably around 20µl or 30µl).
- Transfer the DNA, BSA, the appropriate buffer and ddH2O into a microcentrifuge tube. Finally, add the enzymes to the solution. N.B. The enzymes should be kept on ice before being added to the digestion.
- Incubate for 60-90min at 37°C.
- Use gel electrophoresis to confirm correct digestion.
- Gel purification can be used to obtain the desired digestion product from the gel.
|1/20 Enzyme 1||1.5µl EcoRI|
|1/20 Enzyme 2||1.5µl PstI|
|1/10 BSA||3µl BSA|
|1/10 Buffer* (x 10)||3µl Buffer 4 (x 10)|
|X/10 DNA solution||20µl DNA (pSB1C3)|
|7-X/10 ddH2O||1µl ddH2O|
|Total: 10/10||Total: 30µl|
- The buffer depends on the restriction enzymes used:
|Enzymes used:||Required buffer:|
|Prefix (insert)||EcoRI & SpeI||Buffer 2|
|Suffix (vector)||EcoRI & XbaI||EcoRI buffer|
|Enzymes used:||Required buffer:|
|Prefix (vector)||SpeI & PstI||Buffer 2|
|Suffix (insert)||XbaI & PstI||Buffer3|
A typical ligation reaction mixture is around 10 μl and contains
- 1 μl DNA T4 ligase
- 1 μl DNA T4 ligase buffer (check to ensure it contains ATP) (10x)
- Purified, linearised vector*
- Purified, linearised insert*
- There should be a ratio of 6:1 for moles of insert to vector. This can be calculated using the following equation:
Insert mass (ng) = 6 x (Insert length (bp)/vector length (bp) x Vector mass (ng) Once the solution is made up, the tubes are vortexed and then spun down for around 10 seconds in a microcentrifuge. The ligation is done at 14°C in a water bath in the cold cabinet, and is left overnight.
E. coli Transformation
- One 15ml tube for each sample, in addition to one for a negative control, is put on ice.
- Tubes containing 1ml LB were incubated in a waterbath is set to 42°C .
- Between 25µl and 40µl of competent cells is transferred to each tube.
- The cells are left on ice for 10min.
- 5µl of the DNA sample are transferred into each tube, but ddH2O is added to the control tube(s). The liquids are added directly into the cell culture.
N.B. During pipetting the sides of the tube should not be touched to avoid contamination. Bubbles should be avoided because they can cause the cells stress.
- The tubes are transferred into the 42°C water bath for exactly 45 seconds and then put on ice for 2 minutes. Timing must be exact.
- The tubes are put on a rack and 1ml LB is added to all of them. This levels the temperature of the solution at about 37°C.
- The tubes are then put into a shaking incubator at 37°C for 1 hour.
- The solution from the 15ml tubes is then transferred to a microcentrifuge tube and spun at 13500 rpm for a few seconds.
- The supernatant is discarded and the remaining LB is mixed with the pelleted cells. This increases the concentration of the cells in the LB.
- 50 – 100μl of this solution is then pipetted onto chloraphenicol plates and left overnight at 37°C. The next day colony PCR can be used to examine if the transformation was successful.
Single Colony PCR (SCP)
A master mix is generally used for SCP, as well as Taq polymerase because the high error rate is not an issue here as it is purely confirmatory. Cells from an individual colony are first spread onto a replica plate, and the same loop is then used to inoculate a microcentrifuge tube containing 100μ ddH20 which will later be heated to 95°C to be used in the SCP (the same loop is finally used to inoculate LB for the overnight cultures). The protocol for the first SCP was as follows:
- 19.75μl ddH20
- 2.5μl Barnes buffer
- 1μl template (this comes from the tube that contains 100μ ddH20 and was inoculated with cells.
- 0.5μl dNTPs
- 0.5μl forward primer
- 0.5μl reverse primer
- 0.25μl Taq polymerase.
We also used a positive control (other DNA to which the primers will not anneal) and a negative control (ddH20).
The temperature cycle was as follows:
- 95°C for 30 seconds
- 30 cycles of:
95°C for 30 seconds
62°C for 90 seconds
68 °C for 30 seconds
- 68°C for 10 minutes
- Hold at 4°C
Tubes containing 5ml of LB medium are inoculated with cells from one colony and then 5μl of antibiotic (for example chloramphenicol) is added. They are then left at 37°C overnight.
SDS-PAGE – Protocol
Short for: Sodium Dodecyl Sulfate Polyacrylamide Gel Electrophoresis
- Prepare gel
Two glas plates are cleaned with ethanol and are fitted into a holder. The separation layer of the gel is prepared first, following the recipe but before and after addition of the last two substances the solution should be inverted. The mixture, that now starts to polymerize, in now pipetted between the glass plates until it reaches the green bar. Around 700µl of ethanol are than added ontop of the gel, which is left to solidify. The separating gel contains 10% acrylamide (toxic!) that has been polymerized by TEMED. Stacking gel contains less acryamide for wide pores. After the gel has solidified take out the comb. Once solidified the stacking gel can be prepared using a different recipe but same method. Once the ethanol has been removes the solution is poured onto of the gel and the comb inserted. The gel is now left to solidify.
- Load samples
The proteins, which have been denatured by SDS, are loaded into the wells.
- Run gels
The gel is run at 100V for around 2-3 hours until dye front has reached the bottom of the gel.
- Analysis of results
The gel can be analyzed by staining with Coomassie blue (see protocol) or Western Blot (see protocol)
Other useful information
Using the E.Z.N.A.® Cycle Pure Kit (Omega bio-tek) (ddH2O instead of Elusion Buffer used in last step)
Using the QIAquick® Gel Extraction Kit (250) (ddH2O instead of Elusion Buffer used in last step)
E.Z.N.A.® kit is used.
The QIAGEN HiSpeed Plasmid Midi Kit is used.
- Catechol assay is performed in the plate reader on a 96 well plate
- Each well must be filled with 100um of solution
- Usually use 90ul of cell culture and 10um of catechol solution
- Catechol stock solution is at 100mM concentration. And when added to the well we have a 10fold dilution. For example if an aliquot concentration of 1mM catechol is made, which would be used for assay, when the well is added catechol drops to a concentration of 0.1mM.
- Always dilute catechol with H2O.
- Always have a blank of 90ul medium (the one which you grew the cells overnight) with 10ul catechol solution
- Always have a negative of 90ul growing cells and 10ul of H2O.