MIT iGEM T4 and Quick Ligation

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Calculating Insert Amount

For all ligation reactions, optimal insert:vector molar ratio is 6:1. The appropriate mass ratio for the ligation can be calculated using the following formulas:  {Insert\ Mass\ in\ ng} = 6\times\left[\frac{{Insert\ Length\ in\ bp}}{{Vector\ Length\ in\ bp}}\right]\times{Vector\ Mass\ in\ ng}
I've also used 6:1 crude mass ratios (eg. 60ng of insert, 10ng vector), and this has worked OK, though it's probably not optimal ~~Felix moser


One can use either Quick ligase or T4 ligase for these reactions. Quick is faster but more expensive; T4 is time-tested and proven but takes longer. Here's the protocols for both:

Quick Ligase ligation

Materials

  • 2x Quick ligase buffer (in 40µl aliquots; these are 1-time use since freeze-thaw cycles degrade the ATP in the buffer).
  • Quick Ligase from NEB
  • ddH2O
  • Purified, linearized vector (likely in H2O or EB)
  • Purified, linearized insert (likely in H2O or EB)Italic text

Procedure

For 10µl reaction

Larger volumes can be scaled up if needed

  • 5 μL 2X Quick ligase buffer
  • 0.5 µl Quick ligase
  • 6:1 Molar ratio of insert to vector (~10ng vector). Try to keep total DNA concentration <100ng/rxn for optimal efficiency.
  • Add (4.5 - vector and insert volume)μl ddH2O

Method

  1. Add appropriate amount of deionized H2O to sterile PCR tube
  2. Add in appropriate amounts of vector and insert. Heat the mixture to 42*C for 2min to free up sticky ends (can set up a thermocycler for this).
  3. Add 5 μL of 2X ligation buffer to the tube.
    Pipette buffer up and down before pipetting to ensure that it is well-mixed.
  4. Add 0.5 μL of Quick ligase. PIPETTE half the volume of the mixture UP AND DOWN to ENSURE MIXING OF THE ENZYME.
    Also, the ligase, like most enzymes, is in some percentage of glycerol which tends to stick to the sides of your tip. Just touch your tip to the surface of the liquid when pipetting to ensure accurate volume transfer.
  5. Let the 10 μL solution incubate at room temp for 5-10min.
  6. Denature the ligase at 65°C for 10min.
  7. Store at -20°C





T4 ligase ligation

Materials

  • T4 DNA Ligase
  • 10x T4 DNA Ligase Buffer --> make sure it smells bad (like "wet dog"); if it doesn't smell, it might be bad.
  • Deionized, sterile H2O
  • Purified, linearized vector (likely in H2O or EB)
  • Purified, linearized insert (likely in H2O or EB)Italic text

Procedure

10μl Ligation Mix

Larger ligation mixes are also commonly used

  • 1.0 μL 10X T4 ligase buffer (use 10µl aliquots in -20 freezer; repeated freeze-thaw cycles can degrade the ATP in the buffer that's critical for the ligation rxn)
  • 6:1 Molar ratio of insert to vector (~10ng vector)
  • Add (8.5 - vector and insert volume)μl ddH2O
  • 0.5 μL T4 Ligase

Method

  1. Add appropriate amount of deionized H2O to sterile PCR tube
  2. Add in appropriate amounts of vector and insert. Heat the mixture to 42*C for 2min to free up sticky ends (can set up a thermocycler for this).
  3. Add 1 μL ligation buffer to the tube.
    Pipette buffer up and down before pipetting to ensure that it is well-mixed.
  4. Add 0.5 μL T4 ligase. PIPETTE half the volume of the mixture UP AND DOWN to ENSURE MIXING OF THE ENZYME.
    Also, the ligase, like most enzymes, is in some percentage of glycerol which tends to stick to the sides of your tip. Just touch your tip to the surface of the liquid when pipetting to ensure accurate volume transfer.
  5. Let the 10 μL solution incubate at 16*C for 1hr.
  6. Denature the ligase at 65°C for 10min.
  7. Store at -20°C

Notes

Make sure the buffer is completely melted and dissolved. Precipitate is DTT (or BSA?). Probably best to aliquot this buffer into smaller portions, to reduce the freeze/thaw cycles. In general, make sure the buffer still smells strongly like "wet dog" (Checking if the DTT is still good.)

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