A quick method for cloning fragments with the following ends:

1) a double cut fragment being cloned into a double cut vector if the two restriction sites are non-cohesive.

2) a single cut fragment being cloned into a single cut vector if the vector has been properly dephosphorylated.

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

This method works less well to clone large blunt inserts into a blunt cut vector, and it does not work well to place a sticky ended insert into a sticky ended vector which is not dephosphorylated.

1. Digest vector and insert with enzyme(s), and dephosphorylate the vector if appropreate.

2. Pour a Seaplaque (do not use a substitute!!) low-melt agarose TAE mini-gel with ethidium bromide in the gel and the buffer. Since low-melt agarose gels are fragile take care to make sure that 1) the gel is poured in the cold room so it solidifies quickly enough not to leak out of the mini-gel box 2) Leave enough clearance between comb and gel box such that removing the comb does not rip out the bottom of the well 3) use as low an agarose percentage as possible. I use 0.75% agarose for almost any size fragment except those under 500 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 a low-melt agarose gel .

4) Place gel under the transilluminator and puncture the gel with a 100 µl glass micro-pipette and remove as large a fraction of the vector band as possible. Blow the gel slice into an eppendorf tube, and repeat the procedure for the insert band. See figure on next page. Use a long wave type (360 nm) transulluminator or place a plexiglas shield between the UV box and the gel to protect the DNA from being damaged during theis process.

5. Do not get purify the vector unless there is a need to do so as increasing the agarose concentration in the ligation will reduce the efficiency of the reaction. I standardly do ligations as follows. I do digests with insert only purified in low melt in 30 µl and ligations with both in 50 µl.

Place inserts at 65º C to melt.

3 µl 10X ligation buffer 500 mM Tris pH 7.5, 100 mM MgCl2, 5 mM ATP (added fresh)

50-100 ng vector

H20 to 23 µl

Place at 37ºC

Add 5 µl melted insert (10-200 ng, usually the more the better) using the pipette man to mix quickly. Leave at 37ºC

Add 1/2 µl NEB ligase, mixing well, and move to room temperature. 1-2 hrs is sufficent for most ligations. I sometimes do ligations at 15ºC overnight and this seems to work well.

6. To move the clone into E. coli, I use standard low temp growth comp cell with competent cells (DH5alpha at competency of 108 cfu/µl of pUC18). The transformation procedure it self is very similar to a standard CaCl2 transformation. Place 100 µl of TB buffer in a tube on ice. Melt the ligation mix at 65° C for 5 minutes and add a 5-15 µl aliquot of the ligation mix to the TB solution and vortex quickly. Add 200 µl of competent E. coli to the tube, vortex lightly. Incubate 30-60 minutes on ice, heat shock 90 seconds at 42º C, and return to ice 2 minute. Add 1 ml LB, and incubate 3/4 to 1 hr at 37ºC. I usually plate the whole transformation after spinning down the cells in a low speed centrifuge at 2000 rpm for 5 minutes. If I ligate 100 ng of blunted dephosphorylated vector to 50 ng of a blunt 500 bp PCR in a 30 µl reaction, I'll usually transform 5 ul into cells and plate out the whole tranformation, I normally see approximately 103 colonies (>90% of which are the proper clone).

Ligase Buffer: [660 mM Tris pH 7.5, 66 mM MgCl2, 5 mM ATP (added fresh)]