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Use this if you want amplify a plasmid that cannot be induced. Can be used to amplify most kinds of circular DNA. The kit is intended to amplify circular DNA prior to sequencing but we won’t normally be doing that. The starting point for the protocol described below assumes you are starting from a streaked plate. The manual describes the process for other starting points also.


We begin by denaturing the DNA. We add random primers to this mix, as it cools, the primers can hybridize to the ssDNA. Polymerase is then added to the mixture. It can “roll” around the plasmid many times, starting from everywhere a primer has bound. This leads to massive amplification of the plasmid. The end result is a branched mess of DNA where the plasmid is now in linear form, with many copies concatenated together. A restriction digest can be used to chop up the mix into individual copies of our original plasmid.


  1. A colony of cells containing your circular DNA of choice
  2. Templiphi manual – find this on the side of the refrigerator named – “Marilla”
  3. The Templiphi reagents – find these in the SMUG section of the –80C. They are kept in a small box labeled, surprisingly, Templiphi reagents.
  4. One sterile toothpick for each plasmid you want to use.
  5. Some small PCR tubes.
  6. Ice box to keep Templiphi reagents as cold as possible for as long as possible.


Steps 1-8 can be done in about an hour. The protocol normally requires an overnight step and about 20 mins of work the next morning. At a squeeze, you could get it all done in a day if you have to (shorten the incubation step to 5-6 hours).

  1. Take part of one colony using the toothpick. It’s important that you get as little as possible in order to get the greatest amplification in the end, non-intuitive. Basically, if you have too much of your sample, it will compete with the enzymes you’ll use and you’ll get a lower yield.
  2. Add this part of a colony to 100uL of water in a PCR tube and mix gently with the toothpick.
  3. Do a serial dilution by adding 1uL from this mixture to 4uL of water in a second PCR tube. Add one 1uL of this mixture to another PCR tube containing 4uL of water. I’ll confirm the success of these dilution levels later.
  4. Add 5uL of sample buffer (white cap) to each of the above PCR tubes (we’ll do the experiment at three different dilutions to make sure we get one that works well). You will have to thaw the buffer.
  5. Heat these samples at 95C for 3mins. This denatures the DNA.
  6. Add one 5uL aliquot of reaction buffer (blue cap) to a master mix in a PCR tube for each sample PCR tube you have. Add 0.2uL of enzyme mix to the master mix. You need to use this the same day you make it.
  7. Add 5uL of this mix to each of the sample PCR tubes once they have cooled down.
  8. Put the (labeled) PCR tubes in the 30C room overnight.
  9. To finish the reaction you need to inactivate the enzymes - heat the PCR tubes at 65C for ten minutes.
  10. Allow them to cool to 4C and store as you would the results of a mini-prep.
  11. If you are following the manual protocol, you can ignore the steps referring to cycle sequencing.