Molecular cloning using restriction enzymes

The aim of molecular cloning is to copy (= clone) a piece of DNA (the insert) into a new plasmid backbone. Traditional cloning uses restriction enzymes, which cut DNA at so-called restriction sites, which are DNA sequences that the enzymes recognize. The insert is either copied from genomic or plasmid DNA using PCR or it is cut out from another plasmid using restriction enzymes.


  • Plasmid backbone: at least 3 ug, ideally at a high concentration > 100 ng/ul
  • Template DNA, which is either genomic DNA or plasmid DNA, which the insert will be copied from or cut out from
    • For PCR-based cloning: Primers for amplifying the insert
    • For cutting the insert out of another plasmid: at least 3 ug of the plasmid, ideally at a high concentration > 100 ng/ul
  • Primers for checking the presence of the insert in the final plasmid
  • PCR mix reagents
  • Restriction enzymes and the corresponding buffer(s)
  • Calf Intestinal Polymerase (CIP)
  • T4 DNA ligase and the corresponding buffer


  1. Choose restriction cut-sites and make a map of the desired final plasmid
    Choose the cut-sites with which to cut the plasmid backbone open for the insert. Ideally, cut with two different enzymes rather than one. There is a series of restriction sites at the so-called multiple cloning site (MCS), which Snapgene marks for you as a feature in the plasmid map automatically. Ideally, choose restriction enzymes that have the same optimal temperature and the same optimal buffer so that the two digestions can be run at the same time. Also, make a map of the plasmid you seek to create. Verify, for example, that the additional restriction site that you need for yeast transformation will be present.
  2. For PCR-based cloning: Prepare the insert
    1. Design primers for amplifying the insert
      Choose a pair of primers for amplifying the insert. Add restriction sites, which are compatible with the sites you chose for opening up the plasmid backbone (step 1), to the 3′ ends of the primers. Then add another three random nucleotides to the 3′ ends of the primer sequences so that the restriction enzymes cut the amplified DNA with high efficiency. See link to NEB for more information.
    2. Amplify the DNA insert fragment by PCR
      In order to have enough DNA for cloning, run the reaction in at least 2 tubes with 50 uL PCR mix. Use high-fidelity polymerase (e.g. Phusion) for amplification to avoid mutations in the insert.
    3. Run a small amount of the PCR product on a gel to verify that the PCR reaction produced enough amplified DNA. You should see one band corresponding to the size of the insert you sought to amplify. If there are multiple bands, you should consult someone in the lab about what to do. If the yield is low, you likely have to design other primers for the amplification. Sometimes, it suffices to repeat the PCR reaction with more tubes of the same reaction.
    4. Use the PCR clean-up kit to purify your insert DNA.
    5. Measure the concentration of DNA on Nanodrop. It will be between 20 and 100 ng/ul. If it is below 20 ng/ul it is too low to continue.
  3. Digest the insert and the backbone
    1. Digest the plasmid and the insert, separately, using the restriction enzymes that you chose (see digestion using restriction enzymes protocol). The total volume of the mix should be no more than 50 ul since the reaction mix has to be run on a gel afterwards. If possible, perform a double-digestion by cutting the DNA with the two restriction enzymes at the same time. Pay attention that the concentration of the enzymes should not go above 5% since the glycerol in the enzyme stock inhibits the digestion. Run the digestion for 3 hours at the optimal temperature and in the optimal buffers for the restriction enzymes.
    2. During the last hour of digestion, add Calf Intestinal Alkaline Phosphatase (CIP) to the backbone digestion mix. Cleavage of the phosphate groups at the ends of the linearized backbone prevents self-ligation. Adding CIP is crucial if the backbone is cut with one enzyme only but many of us do not do it with double digestions. Be careful to add CIP only to the plasmid digestion mix and not in the one for the insert. For 10 ug of DNA, add 2 ul of 10.000 U/ml CIP.
    3. Run a gel to separate the DNA fragment of interest from the other fragments. Since the digestion mix is 50 ul, add 10 ul of loading dye (which is 6x concentrated). Use gel combs that create big wells. Use 0.8% agarose gel and run the gel with 120 V for 35 minutes. If you have only used one enzyme to cut the backbone, run a non-cut plasmid together with the one that is cut (the non-cut band should go farther than the cut since the non-linearized plasmid can assume a coiled conformation that has lower resistance when moving through the gel).
    4. Gel extract and purify the DNA. Purify the extracted piece of the gel using the gel extraction kit.
    5. Check the concentration of the purified DNA using the Nanodrop.
  4. Ligate the insert and the backbone (see ligation protocol)
    Use 10 uL as a final volume of the ligation mix. Run a separate negative control with the backbone only. We use different protocols in the lab for the temperature and duration of the ligation. Sahand uses 8 hrs at 4C, then 8 hrs at 16C, then 2 hrs at 25C. No need to purify ligated DNA from enzymes, proceed directly to step 4.
  5. Transform into competent E. coli cells
    Use competent DH5α E. coli cells for the transformation of the ligated product (see bacterial transformation protocol).
  6. Transfer to liquid medium
    1. Pick single colonies that have grown overnight on the LB+antibiotic plate and place them each in 5 ml of liquid LB medium with the appropriate antibiotic. Pick at least 3 different colonies.
    2. Leave the culture on a spinning wheel in the 37°C incubator. The culture should be left overnight but not much longer since the quality of the plasmid DNA declines with time.
  7. Check for the presence of the insert in the ligated plasmid using PCR
    1. Run a PCR with 0.5 ul of bacterial culture to check for the DNA template. Ideally, use a pair of primers where one binds to the insert and the other one binds to the new backbone to confirm not only the presence of the insert but also the expected site of integration and orientation.
    2. Verify the presence and the size of the PCR fragment on a gel. Alternatively, you can digest the new plasmid with restriction enzymes that cut the insert to verify that the cloning worked.
  8. Purify the plasmid using the DNA mini-prep kit
  9. Sequence the region of interest on the plasmid
    Ideally, use a primer that falls into the backbone but produces a sequencing read that mostly contains the insert.


  • Start with a large amount of DNA, which is ideally fresh from a miniprep. Around 10 ug of the plasmid is best since >80% of it is going to be lost during the gel extraction. If you are running a PCR to generate the insert, run between 2 and 8 PCR tubes and purify them all into a 50 uL solution to have enough DNA.
  • During the gel extraction, DNA should be minimally exposed to UV light, as this can cause DNA damage. To further reduce the probability of DNA mutations, use the longer UV wavelength (365 nm, instead of 302 nm) when cutting the gel. Make sure to cut only the part of the gel with the DNA band of interest, not more. While cutting, use the UV protection shield and protect exposed body parts with a lab coat and gloves, as well as eyes and face with a face shield. Cut the gel on a clean and dry surface and pay attention that different pieces of extracted gel do not come in touch with the same surfaces.

Authors: Vojislav Gligorovski and Roxane Dervey
December 2021