Agarose gel electrophoresis

1. Combine 100x agarose powder with 1x TAE buffer in a microwavable flask (eg. 1 g of agarose for 100 mL of TAE). The volume of agarose gel will depend on the size of gel you are making.
2. Microwave for 1-2 min until the agarose is completely dissolved (do not overboil the solution). Stop and swirl the flask every 20 seconds and until the solution is as clear as water.
3. Let agarose solution cool down to about 50°C (when you can comfortably hold the flask with your hand), then add 1 uL of RedSafe to the agarose solution.
4. Seal the ends of a gel tray using masking tape. Pour the agarose into the gel tray with a well comb in place. Let the gel sit at room temperature for 20-30 mins until the gel solidifies.
5. Place the gel into the gel box, fill the gel box with 1 x TAE buffer until the gel is covered then remove the well comb.
6. Mix 2 uL of the digest sample with 3 uL of H2O and 1 uL of 6x loading dye.
7. Load c into the first lane of the gel. Load the remaining digest samples into the gel. Remember to include a negative control (non-digested plasmid).
8. Connect the gel box to a power pack and run the gel at 100V for 1 hr. You should be able to see small bubbles rising in the buffer solution immediately after you turn the power pack on.
9. Carefully take the gel tray to the spectrophotometer and analyse the DNA fragments with UV light. We expect to see a single clear band in digested samples, and a smear for the undigested plasmid at a higher position. Smear and a clear band indicates incomplete digestion.

Gibson Assembly

Materials:

• 5X Isothermal Reaction Mix (6 mL total, Store at -20°C):
• • 3 mL 1 M Tris-Hcl (pH 7.5)
  • 300 μL 1 M MgCl2
  • 60 μL 100 mM dGTP
  • 60 μL 100 mM dATP
  • 60 μL 100 mM dTTP
  • 60 μL 100 mM dCTP
  • 300 μL 1 M DTT
  • 1.5 g PEG-8000
  • 300 μL 100 mM NAD
  • 360 µL water
  • Assembly Master Mix (1.2 mL total, store in 15 µL aliquots at -20°C.):
  • • 320 μL 5X Isothermal Master Mix
    • 0.64 μL 10 U/μL T5 exonuclease
    • 20 μL 2 U/μL Phusion DNA Pol
    • 0.16 μL 40 U/μL T4 DNA Ligase
    • 860 μL water
  
  

  Method:

  1. PCR or digest your fragment of choice and gel purify
  2. If PCR from a methylated DNA template (e.g. propagated plasmid), a DpnI digest can be used to remove the unwanted plasmid. Clean up afterwards.
  3. Thaw a 15 μl assembly mixture aliquot and keep on ice until ready to be used.
  4. Add 5 μl of DNA to be assembled to the master mixture.
  5. The DNA fragments should be in equimolar amounts.
  6. Small fragments (<1 kb) must be added in a five times excess
  7. You can calculate the quantity of each fragment using their molecular weights.
  8. Alternatively, you can use the length of each fragment as a proxy for the molecular weight (assuming similar GC content in all fragments).
  9. Use 10-100 ng of each ~6 kb DNA fragment. For larger DNA segments, increasingly proportionate amounts of DNA should be added (e.g. 250 ng of each 150 kb DNA segment).
  10. Incubate at 50°C for 15 to 60 min (60 min is optimal).

Restriction cloning

1. Set-up the reaction:

   Restriction Enzyme	1µl of each enzyme
   DNA	1 µg
   10X Cutsmart	5 µl (1X)
   Total Reaction Volume	50 µl

2. Incubate for 1 hr at 37°C
3. Heat inactivate at 80°C for 20 minutes

Ligation

1. Set up the following reaction in a microcentrifuge tube on ice:

   2 μl

   Component	20 μl Reaction
   T4 DNA Ligase Buffer (10X)*
   Vector DNA	50 ng
   Insert DNA	A molar ratio of 1:3 vector to insert should be used
   Nuclease-free water	to 20 μl
   T4 DNA Ligase	1 μl

2. Gently mix the reaction by pipetting up and down and microfuge briefly.
3. For cohesive (sticky) ends, incubate at 16°C overnight or room temperature for 10 minutes.
4. Heat inactivate at 65°C for 10 minutes.
5. Chill on ice and transform 1-5 μl of the reaction into 25 μl competent cells.