In Gibson cloning, the ends of the insert do not have to have restriction sites that match corresponding restriction sites in the vector backbone. Instead, the ends of the insert have to have about 30 bps of homology with the DNA flanking the insertion site in the backbone. Because one does not need to create matching restriction site DNA overhangs, Gibson cloning can be easier than restriction cloning. Furthermore, the protocol is faster, and more than two pieces of DNA can be assembled at the same time. Unfortunately, Gibson cloning rarely works with big pieces of DNA (>2 kbs) and can sometimes fail for no discernible reason.
- Plasmid backbone: at least 3 ug, ideally at a high concentration > 100 ng/ul
- Template DNA from which the insert will be copied by PCR
- Primers with tails as explained below for amplifying the insert
- Primers for checking the presence of the insert in the final plasmid
- PCR mix reagents
- Restriction enzymes and the corresponding buffer(s)
- Gibson 2x assembly master mix (NEB)
- Choose restriction cut-sites and make a map of the desired final plasmid
Choose the restriction sites where the plasmid backbone is going to be opened up. It should make no difference whether you open up the plasmid at one or two different sites although it is possible that matching overhangs that are generated with only one restriction enzyme cut are religated without the insert — but we have not tested that. If you have a choice of which restriction enzymes to use, it is best if they have the same optimal temperature and the same optimal buffer, so the digestion by both can be done at the same time. Ideally, make and annotate the map of the expected plasmid before starting. In this way, you can verify whether the cut-sites that are planned to be used for yeast transformation are going to be present in the assembled plasmid.
- Prepare the insert
- Design primers for Gibson Assembly
Each of the two primers for amplifying the insert should contain a 3′ part that anneals to the template DNA with Tm = 56°C and also a 5′ tail that is homologous to the backbone around the cut site. Vojislav recommends at least 20 bps of homology; Sahand recommends 30 bps of homology.
(optional) In Gibson assembly, you can introduce new restriction sites that are going to be unique in the final plasmid.
- Amplify the DNA fragment of interest by PCR using custom Gibson primersIn order to have enough DNA for cloning, run the reaction in at least 2 tubes containing 50 uL PCR mix. Use high-fidelity polymerase (e.g. Phusion) to avoid mutations in the insert.
- Run a gel to test the result of the PCR. If the yield is low, redesign your primers. Sometimes you it suffices to just repeat the PCR with more tubes of the same reaction. Running touch-up PCR (temperature of annealing 56°C in the first 3 cycles and 60°C in all further cycles) might increase the efficiency of the reaction since the binding temperature of the whole Gibson primer is larger once the insert is amplified in the first few rounds of PCR.
- Purify the amplified fragment from the gel using the gel-extraction kit. This step is essential since the PCR product should be separated from any other bands on the gel that might be carried and transformed together with the ligated product (e.g. non-specific PCR bands or the rest of the plasmid in case the plasmid is used as a template).
- Measure the concentration of DNA using the Nanodrop (for purified PCR product it is usually between 20 and 100 ng/ul).
- Design primers for Gibson Assembly
- Prepare the backbone
- Digest the vector backbone using the restriction enzyme(s). The total volume of the mix should be no more than 50 ul since the DNA has to be run on a gel and extracted from the gel after the digestion. Pay attention that the concentration of the restriction enzymes does not exceed 5%, since the glycerol in which enzyme stocks are kept inhibits the digestion. Run the digestion for 3 hours at the temperature and in the buffer defined for the restriction enzymes.
- Run a gel to separate the plasmid backbone from the insert you cut out. 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 a 0.8% agarose gel and run it at 120 V for 35 minutes. If you have used only 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).
- Purify the cut backbone from the gel using a gel-extraction kit.
- Assemble the plasmidMix the Gibson Assembly reaction by following the instructions on the NEB website (https://international.neb.com/products/e2611-gibson-assembly-master-mix#Product%20Information). However, prepare a 10 ul mix instead of the recommended 20 ul mix because the smaller volume suffices.
Example of backbone and insert amounts that have worked in the past:
50 ng of the backbone and three times the amount of insert in mol. (Note: x ng of DNA of length n corresponds to x / n * 1.5 pmol.)
Use half of the reaction mix for assembling the plasmid (5 uL total). Run the negative control (reaction with backbone only). Use a thermocycler to run the Gibson reaction for 1 h at 50°C.
- Transform into competent cells
- Use competent DH5α E. coli cells for the transformation of the assembled product (see bacterial transformation protocol). To avoid having “satellite colonies” of non-resistant bacteria that grow around the ampicillin-resistant colonies, leave the transformed plates at 37°C for no more than 15 h.
- Optional (supposedly helps but has never made a difference for Sahand): To increase the transformation efficiency, grow the transformed cells in 200 uL SOC outgrowth medium for 30 min at 37°C before placing them on solid LB plates that contain the appropriate antibiotic, in our case usually ampicillin.
- Liquid culture transfer
- Pick single colonies that have grown overnight on the plate and place them in 5 ml of liquid LB medium which contains the appropriate antibiotic. Pick no less than 3 colonies.
- Leave bacteria on a spinning wheel in the 37°C incubator to grow overnight. They should be left for around 24 h since the quality of the plasmid DNA starts to drop afterwards.
- Check for the presence of the insert in the assembled plasmid using PCR
- Run a PCR by using 0.5 ul of the liquid bacterial culture for the DNA template. Ideally, use a pair of primers where one binds to the insert and the other one binds to the backbone. Verify the presence and the size of the fragment by running a gel.
- Alternatively, to verify that the cloning worked, you can digest the final plasmid with restriction enzymes that cut the final plasmid but not the original backbone.
- Purify the plasmid using the plasmid mini-prep kit.
- Sequence the region of interest on the plasmid.
Ideally, use a primer that ligates to the backbone but produces a sequencing read that should contain 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