Richard Lab:Amplified insert assembly

From OpenWetWare
Revision as of 21:25, 18 March 2011 by Michael A. Speer (talk | contribs) (Procedure)

Back to Protocols

Amplified Insert Assembly


Amplified Insert Assembly is a method of "BioBricking" two biological parts (i.e. pieces of DNA) together and was developed by Mike Speer and Dr. Tom Richard. For more information on bio-bricking see this link. This method combines the ease and speed of 3A assembly with the reliability of standard assembly. In comparison to 3A assembly, this method can take up to two hours longer; however, the added time spent at the bench is minimal. Major benefits of this assembly method over other bio-brick assembly methods include:

  • no need for gel electrophoresis or gel extraction.
  • the ability to insert small (i.e. invisible on a gel) parts.
  • no need to use multiple antibiotic resistances.
  • no having to make construction vectors.
  • really low background (99% of colonies are correct)
    • This means less sequencing
  • Easy transformation (use homemade competent cells)
  • Less culturing
    • This is because one plasmid prep can supply many PCR inserts. So common parts (i.e. promoters and RBSs) can be used over and over again.

This protocol is typically used to do bio-brick assembly with restriction sites consisting of the following configuration:


Ocassionally other enzymes (e.g. BamHI or HindIII) are used to make protein fusions. See our bio-brick format page for more details.

The two parts you want to assemble will be labeled "insert" and "vector" and will be initially contained on separate plasmids. The eventual goal of assembly is to get these parts on the same plasmid next to one another.



  • Pipettors
  • Microcentrifuge
  • Water baths
  • Thermocycler
  • Electroporator
  • Selective media plates


  • EcoRI Restriction Endonuclease
  • XbaI Restriction Endonuclease
  • SpeI Restriction Endonuclease
  • PstI Restriction Endonuclease
  • DpnI Restriction Endonuclease
  • Vent DNA Polymerase (or another equivalent high fidelity polymerase)
  • Antarctic Phosphatase
  • T4 Ligase


1. Miniprep both "insert" and "vector" from their respective cultures using a kit or this protocol (30 mins).
2. PCR the "insert" plasmid (This will take about 2 hrs, but start the vector digest right away while the insert PCR is cycling).

  • Use a high-fidelity polymerase (e.g. pfu Turbo or Vent).
  • Use the same primers you use for colony PCR (Annealing Temp of 55-60°C).
  • Only run 25-30 cycles as this will help ensure high fidelity.

3. Digest the "vector" for 2 hours. 4. Purify the PCR product using a kit or this protocol. 5. Digest insert for 1 hour (adding DpnI along with the other restriction endonucleases). 6. Add 1μL Antarctic Phosphatase and 6μL AP Buffer to the "vector" digest and incubate until the "insert" digest is done. 7. Kill all reactions by incubating for 20 mins at 80°C. 8. Ligate at a molar ratio of 4:1 (insert:vector). 9. Transform. 10. Plate on plates with the same antibiotic as the "vector" resistance. 11. Celebrate.

  • If you already have PCR insert ready to go (i.e. you ran the PCR the night before from old miniprep) then it only takes about 4 hours.


  • The DpnI eliminates any background from the insert PCR.
  • The phosphatase eliminates any background vector.
  • The "vector" will be digested for a total of thee hours (including nearly one hour with Antarctic Phosphatase)
  • The "insert" will only be digested for one hour. This is okay as there is a lot of it.
  • Detractors of this method may say that it's risky to PCR the inserts because of mutations. We say:
  1. This hasn't been a problem for us.
  2. This is why we use a high-fidelity polymerase
  3. We're sequencing the constructs anyway so we'd spot any mutations.


or instead, discuss this protocol.

Back to Protocols