A quick method for cloning fragments with the following ends (see Note 1):

1) A double cut fragment being cloned into a double cut vector if the two restriction sites are non-cohesive. This is the method we use for most of our cloning experiments. Sticky to blunt also works.

2) A single cut fragment being cloned into a single cut vector if the vector has been properly dephosphorylated. Dephosphorylation is not essential, but will greatly reduce screening needed to identify insert clones (unless some screen or selection is available- e.g. blue/white). We recommend using NEB antiarctic phosphatase which can easily be inactivated at 65°C.

3) A small blunt fragment, such as PCR product, being cloned into a dephosphorylated blunt vector.

1. Digest vector and insert with enzyme(s), and dephosphorylate the vector if appropreate. PCR products should be cleaned up to remove the Taq polymerase before being digested. We recommend running out the inserts in the gel even if they represent the only product in the digest (Note 2).

2. Pour a low-melt agarose (Note 3) TAE mini-gel with ethidium bromide in the gel and the buffer. Since low-melt agarose gels are fragile take care to leave enough clearance between comb and gel box such that removing the comb does not rip out the bottom of the well. We pour these gels in the cold room to help the gel solidify. We also use as low an agarose percentage as possible. We typically use 0.8% agarose for almost any size fragment except those under 300 bp.

3) Load the wells with enough of the digest to have nicely visible bands of vector and insert (i.e ~100 ng of each). Run the gel at 50-100 volts until vector and insert bands are well seperated from other bands of the digests. The gel can be run in the cold room if you have difficulty in handling the gel.

4) Place the gel under a transilluminator and puncture the gel with a 100 µl glass micro-pipette and remove as large a fraction of the vector band and insert bands as possible (Note 4). Blow the gel slice into an eppendorf tube, and repeat the procedure for the insert band. Minimizing DNA damage is key to success of the protocol. We use a medium wavelength (300 nm) transilluminator and keep our gels on a thin glass plate (a 5cm x 8 cm slide) to reduce low wave length UV transmission. Alternatively, use a long wavelength type (360 nm) transilluminator, or place a plexiglas sheet between the UV box and the gel to protect the DNA from being damaged during this process. Our rule of thumb is if you can see it, you can clone it. Typically the punch will be about 8-10 ul.

5. Place vector and inserts at 65º C for 2-3 minutes to melt.

6. Set up ligations in the following order (Note 5)

2.5 ul 10X NEB ligation ligation buffer (Note 6)

14-20 ul H20

1-4 ul melted vector band (20-50 ng)- pipette up and down to mix well

1-4 ul melted insert band (20-50 ng)- pipette up and down to mix well

0.5 ul NEB T4 DNA Ligase (400,000 units /ul)- pipette up and down to mix well

Ligate 1-2 hrs at 15°C for most ligations. I sometimes do ligations at 15ºC overnight for blunt end ligations and this seems to work well.

7. To move the clone into E. coli, I use standard low temp growth competent cells (DH5alpha at competency of 108 cfu/µl of pUC18). Place 100-200 µl of competent E. coli into a cold ependorf tube. Melt the ligation mix at 65° C for 5 minutes and add a 1-5 µl aliquot of the ligation mix to the competent cells and vortex quickly. Incubate 15 minutes on ice, heat shock 40 seconds at 42º C, and return to ice 2 minutes. Add 1 ml SOC (note 7), and incubate 1 hr at 37ºC. I usually plate the 150 ul and keep the rest in the fridge in case I need additional colonies. Alternatively, one can plate the whole transformation after spinning down the cells in a low speed centrifuge at 2000 rpm for 5 minutes.

8. Typically, for a double sticky cloning, we will get hundreds to thousands of colonies on our vector plus insert experimental sample, and less than 5 % as many colonies on our vector only control if the double digest went to completion.

Notes

1. This method may be less efficient for very large blunt inserts and inserting into religatible vectors because the agar reduces diffusion.

2. We used to avoid gel purify vector unless this was required, but we have found that gel purifying the vector often reduces background.

3. We have had great and lasting success using SeaPlaque low melt agarose. I personally would not use a substitute.

4.Punching the bands takes a little practice. Try punching out marker lanes until you get the hang of it.

5. Control ligations could include vector alone or insert alone ligations.

6. We used to make our own ligation buffer [660 mM Tris pH 7.5, 66 mM MgCl2, 5 mM ATP (added fresh)]. NEB ligation buffer works great, but I would aliquot it into 100 ml aliquots so the buffer doesn't go through too many freeze thaw cycles.

7. We used to rescue with LB, but we have found using SOC increases transformation efficiency by at least 10X with our comp cells. They must like the sugar. SOC per liter :20g Bacto Tryptone, 5g Bacto Yeast Extract, 2ml 5M NaCl, 2.5ml of 1M KCl. Sterilize and supplement with 10ml of 1M sterile MgCl2,10ml of 1M sterile MgSO4, 20ml of tterile 1M glucose. We make 5ml aliquots and freeze to reduce odds of contamination.

Updated 3-2-2009